Part I
Foundations of Biochemistry

Facing page: Supernova SN 1987a (the bright “star” at the lower right) resulted
from the explosion of a blue supergiant star in the Large Magellanic Cloud, a galaxy
near the Milky Way. Energy released by nuclear explosions in such supernovae
brought about the fusion of simple atomic nuclei, forming the more complex elements
of which the earth, its atmosphere, and all living things are composed.

Fifteen to twenty billion years ago the universe arose with a cataclysmic explosion that hurled hot, energy-rich subatomic particles into all space. Within seconds, the simplest elements (hydrogen and helium) were formed. As the universe expanded and cooled, galaxies condensed under the influence of gravity. Within these galaxies, enormous stars formed and later exploded as supernovae, releasing the energy needed to fuse simpler atomic nuclei into the more complex elements. Thus were produced, over billions of years, the chemical elements found on earth today. Biochemistry asks how the thousands of different biomolecules formed from these elements interact with each other to confer the remarkable properties of living organisms.

In Part I we will summarize the biological and chemical background to biochemistry. Living organisms operate within the same physical laws that apply to all natural processes, and we begin by discussing those laws and several axioms that flow from them (Chapter 1). These axioms make up the molecular logic of life. They define the means by which cells transform energy to accomplish work, catalyze the chemical transformations that typify them, assemble molecules of great complexity from simpler subunits, form supramolecular complexes that are the machinery of life, and store and pass on the instructions for the assembly of all future generations of organisms from simple, nonliving precursors.

Cells, the units of all living organisms, share certain features; but the cells of different organisms, and the various cell types within a single organism, are remarkably diverse in structure and function. Chapter 2 is a brief description of the common features and the diverse specializations of cells, and of the evolutionary processes that lead to such diversity.

Nearly all of the organic compounds from which living organisms are constructed are products of biological activity. These biomolecules were selected during the course of biological evolution for their fitness in performing specific biochemical and cellular functions. The biomolecules can be characterized and understood in the same terms that apply to the molecules of inanimate matter: the types of bonds between atoms, the factors that contribute to bond formation and bond strength, the three-dimensional structure of molecules, and chemical reactivities. Three-dimensional structure is especially important in biochemistry; the specificity of biological interactions, such as those between enzyme and substrate, antibody and antigen, hormone and receptor, is achieved by close steric complementarity between molecules. Prominent among the forces that stabilize three-dimensional

structure are noncovalent interactions, individually weak but with significant cumulative effects on the structure of biological macromolecules. Chapter 3 provides the chemical basis for later discussions of the structure, catalysis, and metabolic interconversions of individual classes of biomolecules.

Water is the medium in which the first cells arose, and the solvent in which most biochemical transformations occur. The properties of water have shaped the course of evolution and exert a decisive influence on the structure of biomolecules in aqueous solution. Many of the weak interactions within and between biomolecules are strongly affected by the solvent properties of water. Even water-insoluble components of cells, such as membrane lipids, interact with each other in ways dictated by the polar properties of water. In Chapter 4 we consider the properties of water, the weak noncovalent interactions that occur in aqueous solutions of biomolecules, and the ionization of water and of solutes in aqueous solution.

These initial chapters are intended to provide a chemical backdrop for the later discussions of biochemical structures and reactions, so that whatever your background in chemistry or biology, you can immediately begin to follow, and to enjoy, the action.
Part II
Structure and Catalysis

Facing page: End-on view of the triple-stranded collagen superhelix. Collagen,
a component of connective tissue, provides tensile strength and resiliency. Its
strength is derived in part from the three tightly wrapped identical helical strands
(shown in gray, purple, and blue), much the way a length of rope is stronger than
its constituent fibers. The tight wrapping is made possible by the presence of glycine,
shown in red, at every third position along each strand, where the strands are in
contact. Glycine’s small size allows for very close contact.

In Part I we contrasted the complex structure and function of living cells with the relative simplicity of the monomeric units from which the enzymes, supramolecular complexes, and organelles of the cells are constructed. Part II is devoted to the structure and function of the major classes of cellular constituents: amino acids and proteins (Chapters 5 through 8), fatty acids, lipids, and membranes (Chapters 9 and 10), sugars and polysaccharides (Chapter 11), and nucleotides and nucleic acids (Chapter 12). We begin in each case by considering the covalent structure of the simple subunits (amino acids, fatty acids, monosaccharides, and nucleotides). These subunits are a major part of the language of biochemistry; familiarity with them is a prerequisite for understanding more advanced topics covered in this book, as well as the rapidly growing and exciting literature of biochemistry.

After describing the covalent chemistry of the monomeric units, we consider the structure of the macromolecules and supramolecular complexes derived from them. An overriding theme is that the polymeric macromolecules in living systems, though large, are highly ordered chemical entities, with specific sequences of monomeric subunits giving rise to discrete structures and functions. This fundamental theme can be broken down into three interrelated principles: (1) the unique structure of each macromolecule determines its function; (2) noncovalent interactions play a critical role in the structure and function of macromolecules; and (3) the specific sequences of monomeric subunits in polymeric macromolecules contain the information upon which the ordered living state depends. Each of these principles deserves further comment.

The relationship between structure and function is especially evident in proteins, which exhibit an extraordinary diversity of functions. One particular polymeric sequence of amino acids produces a strong, fibrous structure found in hair and wool; another produces a protein that transports oxygen in the blood. Similarly, the special functions of lipids, polysaccharides, and nucleic acids can be understood as a direct manifestation of their chemical structure, with their characteristic monomeric subunits linked in precise functional groups or polymers. Lipids aggregate to form membranes; sugars linked together become energy stores and structural fibers; nucleotides in a polymer become the blueprint for an entire organism.

As we move from monomeric units to larger and larger polymers, the chemical focus shifts from covalent bonds to noncovalent interactions. The covalent nature of monomeric units, and of the bonds that connect them in polymers, places strong constraints upon the shapes

assumed by large molecules. It is the numerous noncovalent interactions, however, that dictate the stable native conformation and provide the flexibility necessary for the biological function of these large molecules. We will see that noncovalent interactions are essential to the catalytic power of enzymes, the arrangement and properties of lipids in a membrane, and the critical interaction of complementary base pairs in nucleic acids.

The principle that sequences of monomeric subunits are information-rich emerges fully in the discussion of nucleic acids in Chapter 12. However, proteins and some polysaccharides are also information-rich molecules. The amino acid sequence is a form of information that directs the folding of the protein into its unique three-dimensional structure, and ultimately determines the function of the protein. Some polysaccharides also have unique sequences and three-dimensional structures that can be recognized by other macromolecules.

For each class of molecules we find a similar structural hierarchy, in which subunits of fixed structure are connected by bonds of limited flexibility, to form macromolecules with three-dimensional structures determined by noncovalent interactions. Together, the molecules described in Part II are the “stuff” of life. We begin with the amino acids.
Part III
Bioenergetics and Metabolism

Facing page: The active site of glyceraldehyde-3-phosphate dehydrogenase,
with the bound cofactor nicotinamide adenine dinucleotide (NAD) shown in red.
This enzyme catalyzes the oxidation of glyceraldehyde-3-phosphate to
1,3-bisphosphoglycerate, a step in glycolysis, a central pathway in glucose metabolism.
This is the earliest known example of an enzymatic reaction in which the energy
released by electron transfer (oxidation) drives the formation of a high-energy
phosphate compound.

Metabolism is a highly coordinated and directed cell activity, in which many multienzyme systems cooperate to accomplish four functions: (1) to obtain chemical energy by capturing solar energy or by degrading energy-rich nutrients from the environment, (2) to convert nutrient molecules into the cell’s own characteristic molecules, including macromolecular precursors, (3) to polymerize monomeric precursors into proteins, nucleic acids, lipids, polysaccharides, and other cell components, and (4) to synthesize and degrade biomolecules required in specialized cellular functions.

Although metabolism embraces hundreds of different enzyme-catalyzed reactions, the central metabolic pathways – our major concern – are few in number and are remarkably similar in all forms of life. Living organisms can be divided into two large groups according to the chemical form in which they obtain carbon from the environment. Autotrophs (such as photosynthetic bacteria and higher plants) can use carbon dioxide from the atmosphere as their sole source of carbon, from which they construct all their carbon-containing biomolecules (see Fig. 2–4). Some autotrophic organisms, such as cyanobacteria, can also use atmospheric nitrogen to generate all their nitrogenous components. Heterotrophs cannot use atmospheric carbon dioxide and must obtain carbon from their environment in the form of relatively complex organic molecules, such as glucose. The cells of higher animals and most microorganisms are heterotrophic. Autotrophic cells are relatively self-sufficient, whereas heterotrophic cells, with their requirements for carbon in more complex forms, must subsist on the products of other cells.

Many autotrophic organisms are photosynthetic and obtain their energy from sunlight, whereas heterotrophic cells obtain their energy from the degradation of organic nutrients made by autotrophs. In our biosphere, autotrophs and heterotrophs live together in a vast, interdependent cycle in which autotrophic organisms use atmospheric CO2 to build their organic biomolecules, some of them generating oxygen from H2O in the process. Heterotrophs in turn use the organic products of autotrophs as nutrients and return CO2 to the atmosphere. The oxidation reactions that produce CO2 also consume O2, converting it to H2O. Thus carbon, oxygen, and water are constantly cycled between the heterotrophic and autotrophic worlds, solar energy ultimately providing the driving force for this massive process (Fig. 1).

Figure 1  The cycling of carbon dioxide and oxygen between the autotrophic (photosynthetic) and the heterotrophic domains in the biosphere. The flow of mass through this cycle is enormous; about 4 × 1011 metric tons of carbon are turned over in the biosphere annually.
All living organisms also require a source of nitrogen, which is necessary for the synthesis of amino acids, nucleotides, and other compounds. Plants are generally able to use either ammonia or soluble nitrates as their sole source of nitrogen, but vertebrate animals must obtain some nitrogen in the form of amino acids or other organic compounds. Only a few organisms – the cyanobacteria and a few species of soil bacteria that live symbiotically on the roots of certain plants (legumes) – are capable of converting (“fixing”) atmospheric nitrogen (N2) into ammonia. Other microbial organisms (nitrifying bacteria) carry out the oxidation of ammonia to nitrites and nitrates. Thus, in addition to the global carbon and oxygen cycle (Fig. 1), a nitrogen cycle operates in the biosphere in which huge amounts of nitrogen undergo cycling and turnover (Fig. 2). The cycling of carbon, oxygen, and nitrogen, which involves many species of living organisms, depends on a proper balance between the activities of the producers (autotrophs) and consumers (heterotrophs) in our biosphere.

Figure 2  The cycling of nitrogen in the biosphere. Gaseous nitrogen (N2) makes up 80% of our atmosphere.

These great cycles of matter are driven by an enormous flow of energy through the biosphere, which begins with the capture of solar energy by photosynthetic organisms and its use to generate energy-rich carbohydrates and other organic nutrients; these nutrients are then used as energy sources by heterotrophic organisms. In the metabolic processes of each organism participating in these cycles, and in all energy-requiring activities, there is a loss of useful energy (free energy) and an inevitable increase in the amount of unavailable energy as heat and entropy. In contrast to the cycling of matter, therefore, energy flows one-way through the biosphere; useful energy can never be regenerated in living organisms from energy dissipated as heat and entropy. Carbon, oxygen, and nitrogen recycle continuously, but energy is constantly transformed into unusable forms.

Figure 3  Energy relationships between catabolic and anabolic pathways. Catabolic pathways deliver chemical energy in the form of ATP, NADH, and NADPH. These are used in anabolic pathways to convert small precursor molecules into cell macromolecules.
Figure 4  Three types of nonlinear metabolic pathways: (a) converging, catabolic; (b) diverging, anabolic; and (c) a cyclic pathway, in which one of the starting materials (oxaloacetate) is regenerated and reenters the pathway. Acetate, a key metabolic intermediate, can be produced by the breakdown of a variety of fuels (a), can serve as the precursor for the biosynthesis of an array of products (b), or can be consumed in the catabolic pathway known as the citric acid cycle (c).
Metabolism, the sum of all of the chemical transformations that occur in a cell or organism, occurs in a series of enzyme-catalyzed reactions that constitute metabolic pathways. Each of the consecutive steps in such a pathway brings about a small, specific chemical change, usually the removal, transfer, or addition of a specific atom, functional group, or molecule. In this sequence of steps (the pathway), a precursor is converted into a product through a series of metabolic intermediates (metabolites). The term intermediary metabolism is often applied to the combined activities of all of the metabolic pathways that interconvert precursors, metabolites, and products of low molecular weight (not including macromolecules).
Catabolism is the degradative phase of metabolism, in which organic nutrient molecules (carbohydrates, fats, and proteins) are converted into smaller, simpler end products (e.g., lactic acid, CO2, NH3). Catabolic pathways release free energy, some of which is conserved in the formation of ATP and reduced electron carriers (NADH and NADPH). In anabolism, also called biosynthesis, small, simple precursors are built up into larger and more complex molecules, including lipids, polysaccharides, proteins, and nucleic acids. Anabolic reactions require the input of energy, generally in the forms of the free energy of hydrolysis of ATP and the reducing power of NADH and NADPH (Fig. 3).

Metabolic pathways are sometimes linear and sometimes branched, yielding several useful end products from a single precursor or converting several starting materials into a single product. In general, catabolic pathways are convergent and anabolic pathways divergent (Fig. 4). Some pathways are even cyclic: one of the starting components of the pathway is regenerated in the series of reactions that converts another starting component into a product. We shall see examples of each type of pathway in the following chapters.

Most organisms have the enzymatic equipment to carry out both the degradation and the synthesis of certain compounds (fatty acids, for example). The simultaneous synthesis and degradation of fatty acids would be wasteful and is prevented by separately regulating anabolic and catabolic reaction sequences: when one occurs, the other is suppressed. Such regulation could not occur if anabolic and catabolic pathways were catalyzed by the same set of enzymes, operating in one direction for anabolism, the opposite for catabolism. Inhibition of an enzyme involved in catabolism would also inhibit the reaction sequence in the anabolic direction. Catabolic and anabolic pathways that connect the same two end points (a fatty acid and acetate, for example) may employ many of the same enzymes, but invariably at least one of the steps is catalyzed by different enzymes in the catabolic and the anabolic directions, and these enzymes are the sites of separate regulation. It is also common for such paired catabolic and anabolic pathways to occur in different cellular compartments. Fatty acid catabolism, for example, occurs in mitochondria, whereas the synthesis of fatty acids takes place in the cytosol. The concentrations of intermediates, enzymes, and regulators can be maintained at different levels in different compartments, further contributing to the separate regulation of catabolic and anabolic reaction sequences. These devices for separation of anabolic and catabolic processes will be of particular interest in our discussions of metabolism.

Metabolic pathways are regulated at three levels. The first and most immediately responsive form of regulation is through the action

of allosteric enzymes, which are capable of changing their catalytic activity in response to stimulatory or inhibitory modulators (p. 230). We shall meet examples of allosteric regulation throughout the following chapters. Metabolic control is exerted at a second level in higher organisms by hormonal regulation. Hormones are chemical messengers released by one tissue that stimulate or inhibit some process in another tissue. Hormones serve to coordinate the metabolic activities of different tissues, and their actions and effects are generally on a somewhat longer time scale than those of allosteric effectors. The third level of metabolic regulation is control of the rate of a metabolic step by regulating the concentration of its enzyme in the cell. The concentration of an enzyme at any given time is the result of a balance between its rate of synthesis and its rate of degradation, both of which are subject to regulation on a time scale of minutes to hours. The number of metabolic transformations that occur in a typical cell can seem overwhelming to a beginning student. Fortunately, there are recurring patterns in the metabolic pathways that make learning easier. Certain types of reactions occur in many different metabolic pathways but always employ the same coenzyme(s) and the same general mechanism. Many of the coenzymes are derived from vitamins (see Table 8–2), compounds essential in the diets of animals. The coenzymes are critical to the reaction mechanisms in which they participate. Once you have learned the general mechanism of a reaction, including the role of the cofactor, the recurring pattern in a variety of metabolic pathways will be easily recognizable. In the chapters that follow, we will usually discuss the general mechanism for each of these reactions when we first encounter the cofactor in its typical role.

In the first half of Part III we consider the major catabolic pathways by which cells obtain energy from the oxidation of various fuels: first, the central pathways of hexose conversion to triose (Chapter 14) and triose oxidation to carbon dioxide (Chapter 15); then the pathways of fatty acid oxidation (Chapter 16) and amino acid oxidation (Chapter 17). Chapter 18 is the pivotal point of our discussion of metabolism; it concerns chemiosmotic energy coupling, the universal mechanism in which a transmembrane electrochemical potential, produced either by substrate oxidation or by light absorption, drives the synthesis of ATP.

The second half of this part describes the major anabolic pathways by which cells use ATP to produce carbohydrates (Chapter 19), lipids (Chapter 20), and amino acids and nucleotides (Chapter 21) from simpler precursors. Finally, in Chapter 22 we step back from the details of the metabolic pathways and consider how those pathways are regulated and integrated in mammals by hormonal mechanisms.

We begin our study of intermediary metabolism with an introduction to bioenergetics (Chapter 13). But before we begin, a final word. Try not to forget that the myriad reactions described on these pages take place in, and play crucial roles in, living organisms. Ask of each reaction and of each pathway, “What is accomplished for the cell or the organism by this reaction or pathway? How does this pathway interconnect with the other pathways occurring simultaneously in the same cell to produce the energy and products required for cell maintenance and growth? How do the multilayered regulatory mechanisms cooperate to balance metabolic and energetic inputs and outputs, achieving the dynamic steady state of life?” Learned with this perspective, metabolism provides fascinating and revealing insights into life.
Chapter 13
Principles of Bioenergetics

Living cells and organisms must perform work to stay alive, to grow, and to reproduce themselves. The ability to harness energy from various sources and to channel it into biological work is a fundamental property of all living organisms; it must have been acquired very early in the process of cellular evolution. Modern organisms carry out a remarkable variety of energy transductions, conversions of one form of energy to another. They use chemical energy in fuels to bring about the synthesis of complex molecules from simple precursors, producing macromolecules with highly ordered structure. They also convert the chemical energy of various fuels into concentration gradients and electrical gradients, motion, heat, and even, in a few organisms such as fireflies, light. Photosynthetic organisms transduce light energy into all of these other forms of energy.

The chemical mechanisms that underlie biological energy transductions have fascinated and challenged biologists for centuries. Antoine Lavoisier, before he lost his head in the French Revolution, recognized that animals somehow transform chemical fuels (foods) into heat and that this process of respiration is essential to life. He observed that

. . . in general, respiration is nothing but a slow combustion of carbon and hydrogen, which is entirely similar to that which occurs in a lighted lamp or candle, and that, from this point of view, animals that respire are true combustible bodies that burn and consume themselves. . . . One may say that this analogy between combustion and respiration has not escaped the notice of the poets, or rather the philosophers of antiquity, and which they had expounded and interpreted. This fire stolen from heaven, this torch of Prometheus, does not only represent an ingenious and poetic idea, it is a faithful picture of the operations of nature, at least for animals that breathe; one may therefore say, with the ancients, that the torch of life lights itself at the moment the infant breathes for the first time, and it does not extinguish itself except at death.*

* From a memoir by Armand Seguin and Antoine Lavoisier, dated 1789, quoted in Lavoisier, A. (1862) Oeuvres de Lavoisier, Imprimerie Imperiale, Paris.

In this century, biochemical studies have revealed much of the chemistry of energy transductions in living organisms. Biological energy transductions obey the same physical laws that govern all other natural processes. It is therefore essential for a student of biochemistry to understand these laws and the ways in which they apply to the flow of energy in the biosphere. In this chapter we first review the laws of

thermodynamics and the quantitative relationships among free energy, enthalpy, and entropy. We then describe the special role of ATP in biological energy exchanges. Finally, we consider the importance of oxidation–reduction reactions in living cells, the energetics of such electron transfer reactions, and the electron carriers commonly employed as cofactors of the enzymes that catalyze these reactions.

Bioenergetics is the quantitative study of the energy transductions that occur in living cells and of the nature and function of the chemical processes underlying these transductions. Although many of the principles of thermodynamics have been introduced in earlier chapters and may be familiar to you, it is worth reviewing the quantitative aspects of these principles.

Many quantitative observations made by physicists and chemists on the interconversion of different forms of energy led to the formulation, in the nineteenth century, of two fundamental laws of thermodynamics. The first law is the principle of the conservation of energy: in any physical or chemical change, the total amount of energy in the universe remains constant, although the form of the energy may change. The second law of thermodynamics, which can be stated in several forms, says that the universe always tends toward more and more disorder: in all natural processes, the entropy of the universe increases.

Living organisms consist of collections of molecules much more highly organized than the surrounding materials from which they are constructed, and they maintain and produce order, seemingly oblivious to the second law of thermodynamics. Living organisms do not violate the second law; they operate strictly within it. To discuss the application of the second law to biological systems, we must first define those systems and the universe in which they occur. The reacting system is the collection of matter that is undergoing a particular chemical or physical process; it may be an organism, a cell, or two reacting compounds. The reacting system and its surroundings together constitute the universe. Some chemical or physical processes can be made to take place in isolated or closed systems, in which no material or energy is exchanged with the surroundings. Living cells and organisms are open systems, which exchange both material and energy with their surroundings; living systems are never at equilibrium with their surroundings.

We have defined earlier in this text three thermodynamic quantities that describe the energy changes occurring in a chemical reaction. Gibbs free energy (G) expresses the amount of energy capable of doing work during a reaction at constant temperature and pressure (p. 8). When a reaction proceeds with the release of free energy (i.e., when the system changes so as to possess less free energy), the free-energy change, ΔG, has a negative sign and the reaction is said to be exergonic. In endergonic reactions, the system gains free energy and ΔG is positive. Enthalpy, H, is the heat content of the reacting system. It reflects the number and kinds of chemical bonds in the reactants and

products. When a chemical reaction releases heat, it is said to be exothermic; the heat content of the products is less than that of the reactants and ΔH has a negative value. Reacting systems that take up heat from their surroundings are endothermic and have positive values of ΔH (p. 66). Entropy, S, is a quantitative expression for the randomness or disorder in a system (Box 13–1). When the products of a reaction are less complex and more disordered than the reactants, the reaction is said to proceed with a gain in entropy (p. 72). The units of ΔG and ΔH are joules/mole or calories/mole (recall that 1 cal equals 4.18 J); units of entropy are joules/mole•degree Kelvin (J/mol•K) (Table 13–1).

Under the conditions existing in biological systems (at constant temperature and pressure), changes in free energy, enthalpy, and entropy are related to each other quantitatively by the equation


in which ΔG is the change in Gibbs free energy of the reacting system,

ΔH is the change in enthalpy of the system, T is the absolute temperature, and ΔS is the change in entropy of the reacting system. By convention ΔS has a positive sign when entropy increases and ΔH has a negative sign when heat is released by the system to its surroundings. Either of these conditions, which are typical of favorable processes, will tend to make ΔG negative. In fact, ΔG of a spontaneously reacting system is always negative.

The second law of thermodynamics states that the entropy of the universe increases during all chemical and physical processes, but it does not require that the entropy increase take place in the reacting system itself. The order produced within cells as they grow and divide is more than compensated for by the disorder they create in their surroundings in the course of growth and division (Box 13–1, case 2). In short, living organisms preserve their internal order by taking from the surroundings free energy in the form of nutrients or sunlight, and returning to their surroundings an equal amount of energy as heat and entropy.

B O X  13–1
Entropy: The Advantages of Being Disorganized

The term entropy, which literally means “a change within”, was first used in 1851 by Rudolf Clausius, one of the promulgators of the second law. A rigorous quantitative definition of entropy involves statistical and probability considerations. However, its nature can be illustrated qualitatively by three simple examples, each of which shows one aspect of entropy. The key descriptors of entropy are randomness or disorder, manifested in different ways.

Case 1: The Teakettle and the Randomization of Heat

We know that steam generated from boiling water can do useful work. But suppose we turn off the burner under a teakettle full of water at 100 °C (the “system”) in the kitchen (the “surroundings”) and allow it to cool. As it cools, no work will be done, but heat will pass from the teakettle to the surroundings, raising the temperature of the surroundings (the kitchen) by an infinitesimally small amount until complete equilibrium is attained. At this point all parts of the teakettle and the kitchen will be at precisely the same temperature. The free energy that was once concentrated in the teakettle of hot water at 100 °C, potentially capable of doing work, has disappeared. Its equivalent in heat energy is still present in the teakettle + kitchen (i.e., the “universe”) but has become completely randomized throughout. This energy is no longer available to do work because there is no temperature differential within the kitchen. Moreover, the increase in entropy of the kitchen (the surroundings) is irreversible. We know from everyday experience that heat will never spontaneously pass back from the kitchen into the teakettle to raise the temperature of the water to 100 °C again.

Case 2: The Oxidation of Glucose

Entropy is a state or condition not only of energy but also of matter. Aerobic organisms extract free energy from glucose obtained from their surroundings. To extract this energy they oxidize the glucose with molecular oxygen, also obtained from the surroundings. The end products of the oxidative metabolism of glucose are CO2 and H2O, which are returned to the surroundings. In this process the surroundings undergo an increase in entropy, whereas the organism itself remains in a steady state and undergoes no change in its internal order. Although some of the entropy arises from the dissipation of heat, entropy also arises from another kind of disorder, illustrated by the equation for the oxidation of glucose by living organisms, which we can write as

C6H12O6 + 6O2   →   6CO2 + 6H2O

or represent schematically as

The atoms contained in 1 molecule of glucose plus 6 molecules of oxygen, a total of 7 molecules, are more randomly dispersed by the oxidation reaction and are now present in a total of 12 molecules (6CO2 + 6H2O).

Whenever a chemical reaction proceeds so that there is an increase in the number of molecules – or when a solid substance, such as glucose, is converted into liquid or gaseous products, which have more freedom to move or fill space than a solid – there is an increase in molecular disorder and thus an increase in entropy.

Case 3: Information and Entropy

The following short passage from Julius Caesar, Act IV, Scene 23, is spoken by Brutus, when he realizes that he must face Mark Antony’s army. It is an information-rich nonrandom arrangement of 125 letters of the English alphabet:

       There is a tide in the affairs of men,

       Which, taken at the flood, leads on to fortune;
       Omitted, all the voyage of their life
       Is bound in shallows and in miseries.

In addition to what this quotation says overtly, it has many hidden meanings. It not only reflects a complex sequence of events in the play, it also echoes the play’s ideas on conflict, ambition, and the demands of leadership. Permeated with Shakespeare’s understanding of human nature, it is very rich in information.

However, if the 125 letters making up this quotation were allowed to fall into a completely random, chaotic pattern, as shown in the following box, they would have no meaning whatsoever.

In this form the 125 letters would contain little or no information, but would be very rich in entropy. Such considerations have led to the conclusion that information is a form of energy; information has been called “negative entropy”. In fact, the branch of mathematics called information theory, which is basic to the programming logic of computers, is closely related to thermodynamic theory. Living organisms are highly ordered, nonrandom structures, immensely rich in information and thus entropy-poor.

Cells are isothermal systems – they function at essentially constant temperature (and at constant pressure). Heat flow is not a source of energy for cells because heat can do work only as it passes from a zone or object at one temperature to a zone or object at a lower temperature. The energy that cells can and must use is free energy, described by the Gibbs free-energy function G, which allows prediction of the direction of chemical reactions, their exact equilibrium position, and the amount of work they can in theory perform at constant temperature and pressure. Heterotrophic cells acquire free energy from nutrient molecules, and photosynthetic cells acquire it from absorbed solar radiation. Both kinds of cells transform this free energy into ATP and other energy-rich compounds, capable of providing energy for biological work at constant temperature.

* Although joules and kilojoules are the standard units of energy and are used
throughout this text, biochemists sometimes express ΔG°’ values in kilocalories
per mole. We have therefore included values in both kilojoules and kilocalories
in this table and in Table 13–5. To convert kilojoules to kilocalories, divide the
number of kilojoules by 4.184.

The composition of a reacting system (a mixture of chemical reactants and products) will tend to continue changing until equilibrium is reached. At the equilibrium concentration of reactants and products, the rates of the forward and reverse reactions are exactly equal and no further net change occurs in the system. The concentrations of reactants and products at equilibrium define the equilibrium constant (p. 90). In the general reaction aA + bB ⇌ cC + dD, where a, b, c, and d are the number of molecules of A, B, C, and D participating, the equilibrium constant is given by


where [A], [B], [C], and [D] are the molar concentrations of the reaction components at the point of equilibrium.

When a reacting system is not at equilibrium, the tendency to move toward equilibrium represents a driving force, the magnitude of which can be expressed as the free-energy change for the reaction, ΔG. Under standard conditions (298 K (25 °C)), when reactants and products are initially present at 1 M concentrations or, for gases, at partial pressures of 101.3 kPa (1 atm), the force driving the system toward equilibrium is defined as the standard free-energy change, ΔG°. By this definition, the standard state for reactions that involve hydrogen ions is [H+] = 1 M, or pH is 0. Most biochemical reactions occur in well-buffered aqueous solutions near pH 7; both the pH and the concentration of water (55.5 M) are essentially constant. For convenience of calculations, biochemists therefore define a slightly different standard state, in which the concentration of H+ is 10–7 M (pH is 7) and that of water is 55.5 M. Physical constants based on this biochemical standard state are written with a prime (e.g., ΔG°’ and Keq) to distinguish them from the constants used by chemists and physicists. Under this convention, when H2O or H+ are reactants or products, their concentrations are not included in equations such as Equation 13–2, but are instead incorporated into the constants ΔG°’ and Keq.

Just as Keq is a physical constant characteristic for each reaction, so too is ΔG°’ a constant. As we noted in Chapter 8 (p. 204), there is a simple relationship between Keq and ΔG°’:

ΔG°’ = –RT ln Keq
The standard free-energy change of a chemical reaction is simply an alternative mathematical way of expressing its equilibrium constant. Table 13–2 shows the relationship between ΔG°’ and Keq. If the equilibrium constant for a given chemical reaction is 1.0, the standard free-energy change of that reaction is 0.0 (the natural logarithm of 1.0 is zero). If Keq of a reaction is greater than 1.0, its ΔG°’ is negative. If Keq is less than 1.0, ΔG°’ is positive. Because the relationship between ΔG°’ and Keq is exponential, relatively small changes in ΔG°’ correspond to large changes in Keq.

It may be helpful to think of the standard free-energy change in another way. ΔG°’ is the difference between the free-energy content of the products and the free-energy content of the reactants under standard conditions. When ΔG°’ is negative, the products contain less free energy than the reactants. The reaction will therefore proceed spontaneously to form the products under standard conditions, because all chemical reactions tend to go in the direction that results in a decrease in the free energy of the system. A positive value of ΔG°’ means that the products of the reaction contain more free energy than the reactants. The reaction will therefore tend to go in the reverse direction if we start with 1.0 M concentrations of all components. Table 13–3 summarizes these points.

As an example, let us make a simple calculation of the standard free-energy change of the reaction catalyzed by the enzyme phosphoglucomutase:

Glucose-1-phosphate  ⇌  glucose-6-phosphate

Chemical analysis shows that whether we start with, say, 20 mM glucose-1-phosphate (but no glucose-6-phosphate) in the presence of phosphoglucomutase, or with 20 mM glucose-6-phosphate, the final equilibrium mixture in either case will contain 1 mM glucose-1-phosphate and 19 mM glucose-6-phosphate at 25 °C and pH 7.0. (Remember that enzymes do not affect the point of equilibrium of a reaction; they merely hasten its attainment.) From these data we can calculate the equilibrium constant:

[glucose-6-phosphate] 19 mM
Keq  =  
  =  19
[glucose-1-phosphate] 1 mM

From this value of Keq we can calculate the standard free-energy change:

ΔG°’ = –RT ln Keq

  = –(8.315 J/mol ∙ K)(298 K)(ln 19)

  = –7,296 J/mol = –7.3 kJ/mol

Because the standard free-energy change is negative, when the reaction starts with 1.0 M glucose-1-phosphate and 1.0 M glucose-6-phosphate, the conversion of glucose-1-phosphate into glucose-6-phosphate
proceeds with a loss (release) of free energy. For the reverse reaction (the conversion of glucose-6-phosphate to glucose-1-phosphate), ΔG°’ has the same magnitude but the opposite sign.

Table 13–4 gives the standard free-energy changes for several representative chemical reactions. Note that hydrolysis of simple esters, amides, peptides, and glycosides, as well as rearrangements and eliminations, proceed with relatively small standard free-energy changes, whereas hydrolysis of acid anhydrides occurs with relatively large decreases in standard free energy. The oxidation of organic compounds to CO2 and H2O proceeds with especially large decreases in standard free energy. However, standard free-energy changes such as those in Table 13–4 tell how much free energy is available from a reaction under standard conditions. To describe the energy released under the conditions that exist within cells, an expression for the actual free-energy change is essential.

We must be careful to distinguish between two different quantities, the free-energy change, ΔG, and the standard free-energy change, ΔG°’. Each chemical reaction has a characteristic standard free-energy

change, which may be positive, negative, or zero, depending on the equilibrium constant of the reaction. The standard free-energy change tells us in which direction and how far a given reaction will go to reach equilibrium when the initial concentration of each component is 1.0 M, the pH is 7.0, and the temperature is 25 °C. Thus ΔG°’ is a constant: it has a characteristic, unchanging value for a given reaction. But the actual free-energy change, ΔG, of a given chemical reaction is a function of the concentrations and of the temperature actually prevailing during the reaction, which are not necessarily the standard conditions as defined above. Moreover, the ΔG of any reaction proceeding spontaneously toward its equilibrium is always negative, becomes less negative as the reaction proceeds, and is zero at the point of equilibrium, indicating that no more work can be done by the reaction.

ΔG and ΔG°’ for any reaction A + B ⇌ C + D are related by the equation

ΔG = ΔG°’ + RT ln  ──────

in which the terms in red are those actually prevailing in the system under observation. The concentration terms in this equation express the effects commonly called mass action. As an example, let us suppose that the reaction A + B ⇌ C + D is taking place at the standard conditions of temperature (25 °C) and pressure (101.3 kPa) but that the concentrations of A, B, C, and D are not equal and that none of them is present at the standard concentration of 1.0 M. To determine the actual free-energy change, ΔG, that will occur under these nonstandard conditions of concentration as the reaction proceeds from left to right, we simply put in the actual concentrations of A, B, C, and D; the values of R, T, and ΔG°’ are the standard values. ΔG will be negative and will approach zero as the reaction proceeds because the actual concentrations of A and B will be getting smaller and the concentrations of C and D will be getting larger. Notice that when a reaction is at equilibrium, where there is no force driving the reaction in either direction and ΔG is equal to zero, Equation 13–3 reduces to

0 = ΔG°’ + RT ln 


ΔG°’ = –RT ln Keq

the equation that, as we noted above (p. 368), relates the standard free-energy change and the equilibrium constant.

Even a reaction for which ΔG°’ is positive can go in the forward direction, if ΔG is negative. This is possible if the term RT ln ([products]/[reactants]) in Equation 13–3 is negative and has a larger absolute value than ΔG°’. For example, the immediate removal of the products of a reaction can keep the ratio [products]/[reactants] well below 1, giving the term RT ln ([products]/[reactants]) a large, negative value.

ΔG and ΔG°’ are expressions of the maximum amount of free energy that a given reaction can theoretically deliver. This amount of energy could be realized only if there were a perfectly efficient device available to trap or harness it. Given that no such device is available, the amount of work done by the reaction at constant temperature and pressure is always less than the theoretical amount.

It is also essential to understand that some reactions that are thermodynamically

favorable (i.e., for which ΔG is large and negative) nevertheless do not occur at measurable rates. For example, firewood can be converted into CO2 and H2O by combustion in a reaction that is very favorable thermodynamically. Nevertheless, firewood is stable for years, because the activation energy (see Fig. 8–4) for its combustion is higher than that provided by room temperature. If the necessary activation energy is provided (with a lighted match, for example), combustion will begin, converting the wood to the more stable products CO2 and H2O and releasing energy as heat and light.

In living cells, reactions that would be extremely slow if uncatalyzed are caused to occur, not by supplying additional heat but by lowering the activation energy with an enzyme (see Fig. 8–4). The free-energy change ΔG for a reaction is independent of the pathway by which the reaction occurs; it depends only on the nature and concentration of the initial reactants and the final products. An enzyme provides an alternative reaction pathway with a lower activation energy, so that at room temperature a large fraction of the substrate molecules have enough thermal energy to overcome the activation barrier, and the reaction rate increases dramatically. Enzymes cannot change equilibrium constants; but they can and do increase the rate at which a reaction proceeds in the direction dictated by thermodynamics.

In the case of two sequential chemical reactions, A ⇌ B and B ⇌ C, each reaction has its own equilibrium constant and each has its characteristic standard free-energy change, ΔG°’1 and ΔG°’2. As the two reactions are sequential, B cancels out and the overall reaction is A ⇌ C. Reaction A ⇌ C will also have its own equilibrium constant and thus will also have its own standard free-energy change, ΔG°’total. The ΔG°′ values of sequential chemical reactions are additive. For the overall reaction A ⇌ C, ΔG°’total is the algebraic sum of the individual standard free-energy changes, ΔG°’1 and ΔG°’2, of the two separate reactions: ΔG°’total = ΔG°’1 + ΔG°’2 . This principle of bioenergetics explains how a thermodynamically unfavorable (endergonic) reaction can be driven in the forward direction by coupling it to a highly exergonic reaction through a common intermediate. For example, the synthesis of glucose-6-phosphate is the first step in the utilization of glucose by many organisms:

Glucose + Pi  →  glucose-6-phosphate + H2O       ΔG°’ = 13.8 kJ/mol

The positive value of ΔG°’ predicts that under standard conditions the reaction will tend not to proceed spontaneously in the direction written. Another cellular reaction, the hydrolysis of ATP to ADP and Pi, is very exergonic:

ATP + H2O  →  ADP + Pi       ΔG°’ = –30.5 kJ/mol

These two reactions share the common intermediates Pi and H2O and may be expressed as sequential reactions:

(1) Glucose + Pi  →  glucose-6-phosphate + H2O
(2) ATP + H2O  →  ADP + Pi

Sum:   ATP + glucose  →  ADP + glucose-6-phosphate

The overall standard free-energy change is obtained by adding the ΔG°’ values for individual reactions:

ΔG°’ = +13.8 kJ/mol + (–30.5 kJ/mol) = –16.7 kJ/mol
The overall reaction is exergonic. In this case, energy stored in the bonds of ATP is used to drive the synthesis of glucose-6-phosphate, a product whose formation from glucose and phosphate is endergonic. The pathway of glucose-6-phosphate formation by phosphate transfer from ATP is different from reactions (1) and (2) above, but the net result is the same as the sum of the two reactions. In thermodynamic calculations, all that matters is the initial and final states; the route between them is immaterial.

We have said that ΔG°’ is a way of expressing the equilibrium constant for a reaction. For reaction (1) above,

Keq1 =  
  = 3.9 × 10−3 M–1

Notice that H2O is not included in this expression. The equilibrium constant for the hydrolysis of ATP is

Keq2 =  
  = 2 × 105 M

The equilibrium constant for the two coupled reactions is

Keq3 =  

=   (Keq1)(Keq2)
=   (3.9 × 10−3 M–1)(2.0 × 105 M)
=   7.8 × 102

By coupling ATP hydrolysis to glucose-6-phosphate synthesis, the Keq for formation of glucose-6-phosphate has been raised by a factor of about 2 × 105.

This strategy is employed by all living cells in the synthesis of metabolic intermediates and cellular components. Obviously, the strategy only works if compounds such as ATP are continuously available. In the following chapters we consider several of the most important cellular pathways for producing ATP.

Having developed some fundamental principles of energy changes in chemical systems, we can now examine the energy cycle in cells and the special role of ATP in linking catabolism and anabolism (see Fig. 1–13). Heterotrophic cells obtain free energy in a chemical form by the catabolism of nutrient molecules and use that energy to make ATP from ADP and Pi. ATP then donates some of its chemical energy to endergonic processes such as the synthesis of metabolic intermediates and macromolecules from smaller precursors, transport of substances across membranes against concentration gradients, and mechanical motion. This donation of energy from ATP generally involves the covalent participation of ATP in the reaction that is to be driven, with the result that ATP is converted to ADP and Pi or to AMP and 2Pi. We discuss here the chemical basis for the large free-energy changes that accompany hydrolysis of ATP and other high-energy phosphate compounds, and show that most cases of energy donation by ATP involve group transfer, not simple hydrolysis of ATP. To illustrate the range of energy transductions in which ATP provides energy, we consider the synthesis of information-rich macromolecules, the transport of solutes across membranes, and motion produced by muscle contraction.

Figure 13–1  The chemical basis for the large free-energy change associated with ATP hydrolysis. (1) Electrostatic repulsion among the four negative charges on ATP is relieved by charge separation after hydrolysis. (2) Inorganic phosphate (Pi) released by hydrolysis is stabilized by formation of a resonance hybrid (left), in which each of the four P–O bonds has the same degree of double-bond character and the hydrogen ion is not permanently associated with any one of the oxygens. (3) The other direct product of hydrolysis, ADP2−, also immediately ionizes (right), releasing a proton into a medium of very low [H+] (pH 7). A fourth factor (not shown) that favors ATP hydrolysis is the greater degree of solvation (hydration) of the products Pi and ADP relative to ATP, which further stabilizes the products relative to the reactants.
Figure 13–2  Formation of Mg2+ complexes partially shields the negative charges and influences the conformation of the phosphate groups in nucleotides such as ATP and ADP.

Figure 13–1 summarizes the chemical basis for the relatively large, negative, standard free energy of hydrolysis of ATP. The hydrolytic cleavage of the terminal phosphoric acid anhydride (phosphoanhydride) bond in ATP separates off one of the three negatively charged phosphates and thus relieves some of the electrostatic repulsion in ATP; the Pi (HPO42−) released by hydrolysis is stabilized by the formation of several resonance forms not possible in ATP; and ADP2−, the other direct product of hydrolysis, immediately ionizes, releasing H+ into a medium of very low [H+](~10–7 M). The low concentration of the direct products favors, by mass action, the hydrolysis reaction.

Although its hydrolysis is highly exergonic (ΔG°’ = –30.5 kJ/mol), ATP is kinetically stable toward nonenzymatic breakdown at pH 7 because the activation energy for ATP hydrolysis is relatively high. Rapid cleavage of the phosphoric acid anhydride bonds occurs only when catalyzed by an enzyme.

Although the ΔG°’ for ATP hydrolysis is –30.5 kJ/mol under standard conditions, the actual free energy of hydrolysis (ΔG) of ATP in living cells is very different. This is because the concentrations of ATP, ADP, and Pi in living cells are not identical and are much lower than the standard 1.0 M concentrations (Table 13–5). Furthermore, the cytosol contains Mg2+, which binds to ATP and ADP (Fig. 13–2). In most enzymatic reactions that involve ATP as phosphoryl donor, the true substrate is MgATP2− and the relevant ΔG°’ is that for MgATP2− hydrolysis. Box 13–2 shows how ΔG for ATP hydrolysis in the intact erythrocyte can be calculated from the data in Table 13–4. ΔG for ATP hydrolysis in intact cells, usually designated ΔGp, is much more negative than ΔG°’; in most cells ΔGp ranges from –50 to –65 kJ/mol. ΔGp is often called the phosphorylation potential. In the following discussion we use the standard free-energy change for ATP hydrolysis, because this allows convenient comparison with the energetics of other cellular reactions for which the actual free-energy changes within cells are not known with certainty.

* For erythrocytes the concentrations are those of the cytosol (human erythrocytes
lack a nucleus and mitochondria). In the other types of cells the data are for the
entire cell contents, although the cytosol and the mitochondria have very different
concentrations of ADP. Phosphocreatine (PCr) is discussed later in this chapter.
B O X  13–2
The Free Energy of Hydrolysis of ATP within Cells: The Real Cost of Doing Metabolic Business

The standard free energy of hydrolysis of ATP has the value –30.5 kJ/mol. In the cell, however, the concentrations of ATP, ADP, and Pi are not only unequal but are also much lower than the standard 1 M concentrations (see Table 13–5). Moreover, the pH inside cells may differ somewhat from the standard pH of 7.0. Thus the actual free energy of hydrolysis of ATP under intracellular conditions (ΔGp) differs from the standard free-energy change, ΔG°’. We can easily calculate ΔGp. For example, in human erythrocytes the concentrations of ATP, ADP, and Pi are 2.25, 0.25, and 1.65 mM, respectively (Table 13–5). Let us assume for simplicity that the pH is 7.0 and the temperature is 25 °C, the standard pH and temperature. The actual free energy of hydrolysis of ATP in the erythrocyte under these conditions is given by the relationship

ΔG = ΔG°’ + RT ln 

Substituting the appropriate values we obtain

(2.50 × 10−4)(1.65 × 10−3)
ΔG = –30,500 J/mol + (8.315 J/mol•K)(398 K) ln 
2.25 × 10−3

= –30,500 J/mol + (2,480 J/mol) ln  (1.83 × 10−4)
= –30,500 J/mol – 21,300 J/mol
= –51,800 J/mol
= –51.8 kJ/mol

Thus ΔGp, the actual free-energy change for ATP hydrolysis in the intact erythrocyte (–51.8 kJ/mol), is much larger than the standard free-energy change (–30.5 kJ/mol). By the same token, the free energy required to synthesize ATP from ADP and Pi under the conditions prevailing in the erythrocyte would be 51.8 kJ/mol.

Because the concentrations of ATP, ADP, and Pi may differ from one cell type to another (Table 13–5), ΔGp for ATP hydrolysis likewise differs. Moreover, in any given cell ΔGp can vary from time to time, depending on the metabolic conditions in the cell and how they influence the concentrations of ATP, ADP, Pi, and H+ (pH). We can calculate the actual free-energy change for any given metabolic reaction as it occurs in the cell, providing we know the concentrations of all the reactants and products of the reaction and other factors (such as pH, temperature, and the concentration of Mg2+) that may affect the equilibrium constant and thus the free-energy change.

Figure 13–3  Hydrolysis of phosphoenolpyruvate (PEP), catalyzed by pyruvate kinase, is followed by spontaneous tautomerization of the product. Tautomerization is not possible in PEP, and thus the product of hydrolysis is stabilized relative to the reactant. Resonance stabilization of Pi also occurs, as shown in Fig. 13–1.
Figure 13–4  Hydrolysis of 1,3-bisphosphoglycerate. The direct product of hydrolysis is 3-phosphoglyceric acid, with an undissociated carboxylic acid group, but dissociation occurs immediately. This ionization and the resonance structures it makes possible stabilize the product relative to the reactants. Resonance stabilization of Pi further contributes to the free-energy change.
Figure 13–6  Hydrolysis of acetyl-coenzyme A, a thioester with a large, negative, free energy of hydrolysis. Thioesters contain a sulfur atom in the position where an oxygen atom is present in oxygen esters. (The complete structure of coenzyme A is shown in Fig. 12–41.)
Figure 13–7  The free energy of hydrolysis of thioesters is large relative to that of oxygen esters. The products of both types of hydrolysis reactions have about the same free-energy content (G), but the thioester has a higher free-energy content than the oxygen ester. Orbital overlap between the O and C atoms allows resonance stabilization in oxygen esters, but orbital overlap between S and C is poorer and little resonance stabilization occurs.

Phosphoenolpyruvate (Fig. 13–3) contains a phosphate ester bond that can undergo hydrolysis to yield the enol form of pyruvate, which immediately tautomerizes to the more stable keto form. Because the product of hydrolysis can exist in either of two tautomeric forms (enol and keto), whereas the reactant has only one form (enol), the product is stabilized relative to the reactant. This is the main reason for the high standard free energy of hydrolysis of phosphoenolpyruvate: ΔG°’ = –61.9 kJ/mol.

Another three-carbon compound, 1,3-bisphosphoglycerate (Fig. 13–4), contains an anhydride bond between the carboxyl group at C-1 and phosphoric acid. Hydrolysis of this acyl phosphate is accompanied by a large, negative, standard free-energy change (ΔG°’ = –49.3 kJ/mol), which can be rationalized in terms of the structure of reactant and products. When H2O is added across the anhydride bond, one of the direct products (3-phosphoglyceric acid) can immediately lose a proton. Removal of this direct product favors the forward reaction and results in the formation of the carboxylate ion (3-phosphoglycerate), which has two equally probable resonance forms (Fig. 13–4).

In phosphocreatine (Fig. 13–5), the P–N bond can be hydrolyzed to generate free creatine and Pi. As in the previous cases, the release of Pi favors the forward reaction. Creatine can exist in two resonance forms, and this resonance stabilization of the product favors the forward reaction. The standard free-energy change in this reaction is large, about –43 kJ/mol.

Figure 13–5  Hydrolysis of phosphocreatine. Breakage of the P–N bond in phosphocreatine produces creatine, which forms a resonance hybrid and is thus stabilized. The other product, Pi, is also resonance stabilized.
In all of the reactions that liberate Pi, the several resonance forms available to Pi (Fig. 13–1) stabilize this product relative to the reactant, further contributing to a negative free-energy change for the hydrolysis reactions. Table 13–6 lists the standard free energies of hydrolysis for a number of phosphorylated compounds.

Source: Data mostly from Jencks, W.P. (1976) In Handbook of Biochemistry and
Molecular Biology
, 3rd edn (Fasman, G.D., ed), Physical and Chemical Data, Vol. I,
pp. 296–304, CRC Press, Cleveland, OH.

Thioesters are compounds that do not release Pi on hydrolysis but nevertheless have large, negative, standard free energies of hydrolysis. Acetyl-coenzyme A (Fig. 13–6) is a thioester that we will encounter repeatedly in later chapters. There is no resonance stabilization in thioesters comparable to that in oxygen esters (Fig. 13–7); consequently, the difference in free energy between the thioester and its hydrolysis products, which are resonance-stabilized, is greater than that for comparable oxygen esters. In both cases, hydrolysis of the ester generates a carboxylic acid, which can ionize and assume two resonance forms as described above for acyl phosphates. The free energy of hydrolysis for acetyl-CoA is large and negative, about –31 kJ/mol.

To summarize, compounds with large, negative, standard free energies of hydrolysis give products that are more stable than the reactants because of one or more of the following: (1) the bond strain in reactants due to electrostatic repulsion is relieved by charge separation, as in the case of ATP (described earlier), (2) the products are stabilized by ionization, as in the case of ATP, acyl phosphates, and thioesters, (3) the products are stabilized by isomerization (tautomerization), as for phosphoenolpyruvate, and/or (4) the products are stabilized by resonance, as for creatine from phosphocreatine, the carboxylate ion from acyl phosphates and thioesters, and phosphate (Pi) from all of the phosphorylated compounds.
Figure 13–8  The contribution of ATP to a reaction is often shown with a single arrow (a), but is almost always a two-step process, such as that shown here for the reaction catalyzed by ATP-dependent glutamine synthetase (b).
Figure 13–9  Flow of phosphate groups, represented by Ⓟ from high-energy phosphate donors via ATP to acceptor molecules (such as glucose and glycerol) to form their low-energy phosphate derivatives. This flow of phosphate groups, which is catalyzed by enzymes called kinases, proceeds with an overall loss of free energy under intracellular conditions. Hydrolysis of low-energy phosphate compounds releases Pi, which has an even lower group transfer potential.
Figure 13–10  Both phosphoric acid anhydride bonds in ATP are eventually broken in the formation of palmitoyl-coenzyme A. In the first step of the reaction, ATP donates adenylate (AMP), forming the fatty acyl adenylate and releasing PPi. The pyrophosphate is subsequently hydrolyzed by inorganic pyrophosphatase. The “energized” fatty acyl group is then transferred to coenzyme A.

Throughout this book we will refer to reactions or processes for which ATP supplies energy, and the contribution of ATP to these reactions will commonly be indicated as in Figure 13–8a, with a single arrow showing the conversion of ATP into ADP and Pi, or of ATP into AMP and PPi (pyrophosphate). When written this way, these reactions of ATP appear to be simple hydrolysis reactions in which water displaces either Pi or PPi, and one is tempted to say that an ATP-dependent reaction is “driven by the hydrolysis of ATP”. This is not the case. ATP hydrolysis per se usually accomplishes nothing but the liberation of heat, which cannot drive a chemical process in an isothermal system.

Single reaction arrows such as those in Figure 13–8a almost invariably represent two-step processes (Fig. 13–8b) in which part of the ATP molecule, either a phosphoryl group or the adenylate moiety (AMP), is first transferred to a substrate molecule or to an amino acid residue in an enzyme, becoming covalently attached to and raising the free-energy content of the substrate or enzyme. In the second step, the phosphate-containing moiety transferred in the first step is displaced,

generating either Pi or AMP. Thus ATP participates in the enzyme-catalyzed reaction to which it contributes free energy. There is one important class of exceptions to this generalization: those processes in which noncovalent binding of ATP (or of GTP), followed by its hydrolysis to ADP and Pi, provides the energy to cycle a protein between two conformations, producing mechanical motion, as in muscle contraction or in the movement of enzymes along DNA (discussed below).

The phosphate compounds found in living organisms can be divided arbitrarily into two groups, based on their standard free energies of hydrolysis (Fig. 13–9). “High-energy” compounds have a ΔG°’ of hydrolysis more negative than –25 kJ/mol; “low-energy” compounds have a less negative ΔG°’. ATP, for which ΔG°’ of hydrolysis is –30.5 kJ/mol (–7.3 kcal/mol), is a high-energy compound; glucose-6-phosphate, with a standard free energy of hydrolysis of –13.8 kJ/mol (–3.3 kcal/mol), is a low-energy compound.

The term “high-energy phosphate bond”, although long used by biochemists, is incorrect and misleading, as it wrongly suggests that the bond itself contains the energy. In fact, the breaking of chemical bonds requires an input of energy. The free energy released by hydrolysis of phosphate compounds thus does not come from the specific bond that is broken but results from the products of the reaction having a smaller free-energy content than the reactants. For simplicity, we will sometimes use the term “high-energy phosphate compound” when referring to ATP or other phosphate compounds with a large, negative, standard free energy of hydrolysis.

From the additivity of free-energy changes of sequential reactions, one can see that the synthesis of any phosphorylated compound can be accomplished by coupling it to the breakdown of another phosphorylated compound with a more negative free energy of hydrolysis (Fig. 13–9). One can therefore describe phosphorylated compounds as having a high or low phosphate group transfer potential. The phosphate group transfer potential of phosphoenolpyruvate is very high, that of ATP is high, and that of glucose-6-phosphate is low.

Much of catabolism is directed toward the synthesis of high-energy phosphate compounds, but their formation is not an end in itself; it is the means of activating a very wide variety of compounds for further chemical transformation. The transfer of a phosphoryl group to a compound effectively puts free energy into that compound, so that it has more free energy to give up during subsequent metabolic transformations. We described above how the synthesis of glucose-6-phosphate is accomplished by phosphoryl group transfer from ATP. We shall see in the next chapter that this phosphorylation of glucose activates or “primes” the glucose for catabolic reactions that occur in nearly every living cell.

In some reactions that involve ATP, both of its terminal phosphate groups are released in one piece as PPi. Simultaneously, the remainder of the ATP molecule (adenylate) is joined to another compound, which is thereby activated. For example, the first step in the activation of a fatty acid either for energy-yielding oxidation (Chapter 16) or for use in the synthesis of more complex lipids (Chapter 20) is its attachment to the carrier coenzyme A (Fig. 13–10). The direct condensation of a fatty acid with coenzyme A is endergonic, but the formation of fatty acyl–CoA is made exergonic by coupling it to the net breakdown, in two steps, of ATP.

In the first step, adenylate (AMP) is transferred from ATP to the carboxyl group of the fatty acid, forming a mixed anhydride (fatty acyl adenylate) and liberating PPi. In the second step, the thiol group of coenzyme A displaces the adenylate group and forms a thioester with the fatty acid. The sum of these two reactions is the exergonic hydrolysis of ATP to AMP and PPiG°’ = –32.2 kJ/mol) and the endergonic formation of fatty acyl–CoA (ΔG°’ = 31.4 kJ/mol).

The formation of fatty acyl–CoA is made energetically favorable by a third step, in which the PPi formed in the first step is hydrolyzed by the ubiquitous enzyme inorganic pyrophosphatase to yield 2Pi:

PPi + H2O  →  2Pi       ΔG°’ = –33.4 kJ/mol

Thus, in the activation of a fatty acid, both of the phosphoric acid anhydride bonds of ATP are broken. The resulting ΔG°’ is the sum of the ΔG°’ values for the breakage of these bonds:

ATP + 2H2O  →  AMP + 2Pi       ΔG°’ = –65.6 kJ/mol

The activation of amino acids before their polymerization into proteins (Chapter 26) is accomplished by an analogous set of reactions. An aminoacyl adenylate is first formed from the amino acid and ATP, with the elimination of PPi. The adenylate group is then displaced by a transfer RNA, which is thereby joined to the amino acid. In this case, too, the PPi formed in the first step is hydrolyzed by inorganic pyrophosphatase. An unusual use of the cleavage of ATP to AMP and PPi occurs in the firefly, which uses ATP as an energy source to produce light flashes (Box 13–3, p. 382).

The AMP produced in adenylate transfers is returned to the ATP cycle by the action of adenylate kinase, which catalyzes the reversible reaction

ATP + AMP     ADP + ADP       ΔG°’ ≈ 0

The ADP so formed can be phosphorylated to ATP, using reactions described in detail in later chapters.
Figure 13–11  Nucleoside triphosphates are the substrates for RNA synthesis. With each nucleoside monophosphate added to the growing chain, one PPi is released and then hydrolyzed to two Pi. The hydrolysis of two phosphoric acid anhydride bonds for each nucleotide added provides energy for forming the bonds in the RNA polymer and for assembling a specific sequence of nucleotides.

When simple precursors are assembled into high molecular weight polymers with defined sequences (DNA, RNA, proteins), as described in detail in Part IV, energy is required both for the condensation of monomeric units and for the creation of ordered sequences. The precursors for DNA and RNA synthesis are nucleoside triphosphates, and polymerization is accompanied by cleavage of the phosphoric acid anhydride linkage between the α- and β-phosphates, with the release of PPi (Fig. 13–11). The moieties transferred to the growing polymer in these polymerization reactions are adenylate (AMP), guanylate (GMP), cytidylate (CMP), or uridylate (UMP) for RNA synthesis, and their deoxy analogs for DNA synthesis. We have seen that the activation of amino acids for protein synthesis involves the donation of adenylate groups from ATP, and we shall see later that the formation of peptide bonds on the ribosome is also accompanied by GTP hydrolysis (Chapter 26). In all of these cases, the exergonic breakdown of a nucleoside triphosphate is coupled to the endergonic process of synthesizing a polymer of a specific sequence.

B O X  13–3
Firefly Flashes: Glowing Reports of ATP

Figure 1  The firefly, a beetle of the Lampyridae family.

Many fungi, marine microorganisms, jellyfish, and crustaceans as well as the firefly (Fig. 1) are capable of generating bioluminescence, which requires considerable amounts of energy. In the firefly, ATP is used in a set of reactions that converts chemical energy into light energy. From many thousands of firefly lanterns collected by children in and around Baltimore, William McElroy and his colleagues at The Johns Hopkins University isolated the principal biochemical components involved, luciferin (Fig. 2), a complex carboxylic acid, and luciferase, an enzyme. The generation of a light flash requires activation of luciferin by an enzymatic reaction with ATP in which a pyrophosphate cleavage of ATP occurs, to form luciferyl adenylate (Fig. 2).

This compound is then acted upon by molecular oxygen and luciferase to bring about the oxidative decarboxylation of the luciferin to yield oxyluciferin. This reaction, which has intermediate steps, is accompanied by emission of light (Fig. 2). The color of the light flash differs with firefly species and appears to be determined by differences in the structure of the luciferase. Luciferin is then regenerated from oxyluciferin in a subsequent series of reactions. Other bioluminescent organisms use other types of enzymatic reactions to generate light.

In the laboratory, pure firefly luciferin and luciferase are used to measure minute quantities of ATP by the intensity of the light flash produced. As little as a few picomoles (10−12 mol) of ATP can be measured in this way.

Figure 2  Important components in firefly bioluminescence, and the firefly bioluminescence cycle.

ATP can supply the energy for transporting an ion or a molecule across a membrane into another aqueous compartment where its concentration is higher. Recall from Chapter 10 that the free-energy change (ΔGt) for the transport of a nonionic solute from one compartment to another is given by

ΔGt = RT ln (C2/C1)

where C1 is the molar concentration of the solute in the compartment from which the ion or molecule moves and C2 is its molar concentration in the compartment into which it moves. When a proton or other charged species moves across a membrane without a counterion, the separation of electrical charge requires extra electrical work beyond the osmotic work against a concentration gradient. The extra electrical work is Z ℱ Δψ, where Z is the (unitless) electrical charge of the transported species, Δψ is the transmembrane electrical potential (in volts), and ℱ is the Faraday constant (96.48 kJ/V•mol). The total energy cost of moving a charged species against an electrochemical gradient is

ΔGt = RT ln (C2/C1) + Z ℱ Δψ

Transport processes are major consumers of energy; in tissues such as human kidney and brain, as much as two-thirds of the energy consumed at rest is used to pump Na+ and K+ across plasma membranes via the Na+K+ ATPase. Na+ and K+ transport is driven by cyclic phosphorylation and dephosphorylation of the transporter protein, with ATP as the phosphate donor (see Fig. 10–23). Na+-dependent phosphorylation of the Na+K+ ATPase forces a change in the protein’s conformation, and K+-dependent dephosphorylation favors return to the original conformation. Each cycle in the transport process results in the conversion of ATP to ADP and Pi, and it is the free-energy change of ATP hydrolysis that drives the pumping of Na+ and K+. In animal cells, the net hydrolysis of one ATP is accompanied by the outward transport of three Na+ ions and the uptake of two K+ ions.

Figure 13–12  ATP hydrolysis drives the cross-bridge cycle during the sliding motion of actin–myosin complexes in muscle. This proposed mechanism begins with each myosin head bound to an actin filament. Binding of ATP to myosin (a) causes dissociation of the actin–myosin cross-bridge. ATP hydrolysis (b) leaves myosin with bound ADP and Pi, which favors a different conformation of the myosin head. In this conformation, the myosin head binds to an adjacent actin filament (c) when elevated cytosolic Ca2+ signals contraction. This cross-bridge formation induces the release of bound ADP and Pi (d), which provides the free energy for a conformational change in the myosin head; the head tilts, forcing the thin (actin) filament to slide relative to the thick (myosin) flament, producing contraction. ATP then binds to the myosin head to dissociate the cross-bridge and start another cycle. Each cycle occurs in about 1 msec.

In the contractile system of skeletal muscle cells, myosin and actin are specialized to transduce the chemical energy of ATP into motion. ATP binds tightly but noncovalently to the head portion of one conformation of myosin, holding the protein in that conformation. When myosin (which is also an ATPase) catalyzes the hydrolysis of its bound ATP, the ADP and Pi produced dissociate from the protein, allowing it to relax into a second conformation until another molecule of ATP binds (Fig. 13–12). The binding and subsequent hydrolysis of ATP thus provide the energy that forces cyclic changes in the conformation of the myosin head. The change in conformation of many individual myosin molecules results in the sliding of myosin fibrils along actin filaments (see Fig. 7–32), which translates into macroscopic contraction of the muscle fiber.

This production of mechanical motion at the expense of ATP is one of the few cases in which ATP hydrolysis per se, and not group transfer from ATP, is the source of the chemical energy in a coupled process. The energy-dependent reactions catalyzed by helicases, RecA protein, and some topoisomerases (Chapter 24) and by certain GTP-binding proteins (Chapter 22) also involve direct hydrolysis of phosphoric acid anhydride bonds.

The transfer of phosphate groups is one of the central features of metabolism. Metabolic electron transfer reactions are also of crucial importance. These oxidation–reduction reactions involve the loss of electrons by one chemical species, which is thereby oxidized, and the gain by another, which is reduced. The flow of electrons in oxidation–reduction reactions is responsible, directly or indirectly, for all of the work done by living organisms. In nonphotosynthetic organisms, the source of electrons is reduced compounds (food); in photosynthetic organisms, the initial electron donor is a chemical species excited by the absorption of light. The path of electron flow in metabolism is complex. Electrons move from various metabolic intermediates to specialized electron carriers in enzyme-catalyzed reactions. Those carriers in turn donate electrons to acceptors with higher electron affinities, with the release of energy. Cells contain a variety of molecular energy transducers, which convert the energy of electron flow into useful work.

We begin our discussion with a description of the general types of metabolic reactions that involve electron transfers. After considering the theoretical and experimental basis for measuring energy changes in oxidation reactions in terms of electromotive force, we will discuss the relationship between this force, expressed in volts, and the free-energy change, expressed in joules. We conclude by introducing the structures and oxidation–reduction chemistry of the most common of the specialized electron carriers, which we shall meet repeatedly in later chapters.

Figure 13–13  The analogy between macroscopic (a) and microscopic (b) electrical circuits. In both circuits, the energy of electron flow is harnessed to do work.

The conversion of electron flow to biological work requires molecular transducers, analogous to the electric motors that convert electron flow through macroscopic circuits into mechanical motion. The analogy between a circuit connecting a battery with an electric motor and the submicroscopic electron circuits in cells is instructive.

In the macroscopic circuit (Fig. 13–13a), the source of electrons is a battery containing two chemical species that differ in affinity for electrons. The electrical wires provide a pathway for electron flow from the chemical species at one pole of the battery, through the motor, to the chemical species at the other pole of the battery. Because the two chemical species differ in their affinity for electrons, electrons flow spontaneously through the circuit, driven by a force proportional to the difference in electron afimity, the electromotive force. The electromotive force (typically a few volts) can accomplish work if an appropriate energy transducer such as a motor is placed in the circuit. The motor can be coupled to a variety of mechanical devices to accomplish work.

In an analogous biological “circuit” (Fig. 13–13b), the source of electrons is a relatively reduced compound such as glucose. As glucose is enzymatically oxidized, electrons are released and flow spontaneously through a series of electron carrier intermediates to another chemical species with a high affinity for electrons, such as O2. Electron flow is spontaneous and exergonic, because O2 has a higher affinity for electrons than do the intermediates that donate electrons. The resulting electromotive force provides energy to molecular transducers that do biological work. In the mitochondrion, for example, membrane-bound transducers couple electron flow to the production of a transmembrane

pH difference, accomplishing osmotic and electrical work. The proton gradient thus formed has potential energy, sometimes called proton-motive force by analogy with electromotive force. Another molecular transducer in the mitochondrial membrane uses the proton-motive force to do chemical work: ATP is synthesized from ADP and Pi as protons flow spontaneously across the membrane. Similarly, membrane-localized transducers in E. coli convert electromotive to proton-motive force, which is then used to power flagellar motion.

The principles of electrochemistry that govern energy changes in the circuit with a motor and battery apply with equal validity to the microscopic processes accompanying electron flow in living cells. We turn now to a review of those principles.

Although oxidation and reduction must occur together, it is convenient when describing electron transfers to consider the two halves of an oxidation–reduction reaction separately. For example, the oxidation of ferrous ion by cupric ion:

Fe2+ + Cu2+   ⇌   Fe3+ + Cu+

can be described in terms of two half reactions:

(1)   Fe2+   ⇌   Fe3+ + e

(2)   Cu2+ + e   ⇌   Cu+

The electron-donating molecule in an oxidation–reduction reaction is called the reducing agent or reductant; the electron-accepting molecule is the oxidizing agent or oxidant. A given agent, such as an iron cation in the ferrous (Fe2+) and the ferric (Fe3+) state, functions as a conjugate

reductant–oxidant pair (redox pair), just as an acid and corresponding base function as a conjugate acid–base pair. Recall from Chapter 4 that in acid–base reactions we can write the general equation: proton donor ⇌ H+ + proton acceptor. In redox reactions we can write a similar general equation: electron donor ⇌ e + electron acceptor. In the reversible half reaction (1) above, Fe2+ is the electron donor and Fe3+ is the electron acceptor; together, Fe2+ and Fe3+ constitute a conjugate redox pair.

The electron transfers in oxidation–reduction reactions involving organic compounds are not fundamentally different from those that occur with inorganic species. In Chapter 11 we considered the oxidation of a reducing sugar (an aldehyde or ketone) by cupric ion (see Fig. 11–10a):

This overall reaction can be expressed as two half reactions:

Because two electrons are removed from the aldehyde carbon, the second half reaction (the one-electron reduction of cupric to cuprous ion) must be doubled to balance the overall equation.

Figure 13–14  The oxidation states of carbon. Each of the arrows indicates an oxidation reaction; all except the last reaction are oxidations brought about by dehydrogenation.

Carbon occurs in living cells in five different oxidation states (Fig. 13–14). In the most reduced compounds carbon atoms are rich in electrons and in hydrogen, whereas in the more highly oxidized compounds a carbon atom is bonded to more oxygen and to less hydrogen. In the oxidation of ethane to ethanol (Fig. 13–14), the compound does not lose a hydrogen but one of the carbon atoms does; the hydrogen of the –OH group is, of course, not bonded directly to carbon. In the series of compounds shown in Figure 13–14, oxidation of a carbon atom is synonymous with its dehydrogenation. When a carbon atom shares an electron pair with another atom such as oxygen, the sharing is unequal, in favor of the more electronegative atom (oxygen). Thus oxidation has the effect of removing electrons from the carbon atom.

Not all biological oxidation–reduction reactions involve oxygen and carbon. For example, the conversion of molecular nitrogen into ammonia, 6H+ + 6e + N2 → 2NH3, represents a reduction of the nitrogen atoms.

Electrons are transferred from one molecule to another in one of four different ways:

 1. They may be transferred directly as electrons. For example, the Fe2+/Fe3+ redox pair can transfer an electron to the Cu+/Cu2+ redox pair:

Fe2+ + Cu2+   ⇌   Fe3+ + Cu+

2. Electrons may be transferred in the form of hydrogen atoms. Recall that a hydrogen atom consists of a proton (H+) and a single electron (e). In this case we can write the general equation

AH2   ⇌   A + 2e + 2H+
 where AH2 acts as the hydrogen (or electron) donor. AH2 and A together constitute a conjugate redox pair, which can reduce another compound B by transfer of hydrogen atoms:

AH2 + B   ⇌   A + BH2

3. Electrons may be transferred from an electron donor to an acceptor in the form of a hydride ion ( ⁚ H), which includes two electrons, as in the case of NAD-linked dehydrogenases described below.

4. Electron transfer also takes place when there is a direct combination of an organic reductant with oxygen, to give a product in which the oxygen is covalently incorporated, as in the oxidation of a hydrocarbon to an alcohol:

R–CH3 + ½O2   →   R–CH2–OH

 In this reaction the hydrocarbon is the electron donor and the oxygen atom is the electron acceptor.

All four types of electron transfer occur in cells. The neutral term reducing equivalent is commonly used to designate a single electron equivalent participating in an oxidation–reduction reaction, no matter whether this equivalent be in the form of an electron per se, a hydrogen atom, or a hydride ion, or whether the electron transfer takes place in a reaction with oxygen to yield an oxygenated product. Because biological fuel molecules usually undergo enzymatic dehydrogenation to lose two reducing equivalents at a time, and because each oxygen atom can accept two reducing equivalents, biochemists by convention refer to the unit of biological oxidations as two reducing equivalents passing from substrate to oxygen.
Figure 13–15  Measurement of the standard reduction potential (E0′) of a redox pair. Electrons flow from the test electrode to the reference electrode, or vice versa. The ultimate reference half-cell is the hydrogen electrode, as shown here. The arbitrary electromotive force (emf) of this electrode is 0.00 V. At pH 7, E0′ for the hydrogen electrode is –0.414 V. The direction of electron flow depends upon the relative electron “pressure” or potential of the two cells. A salt bridge containing a saturated KCl solution provides a path for counter-ion movement between the test cell and the reference cell. From the observed emf and the known emf of the reference cell, the emf of the test cell containing the redox pair is obtained. The cell that gains electrons has, by convention, the more positive reduction notential.

When two conjugate redox pairs are present together in solution, electron transfer from the electron donor of one pair to the electron acceptor of the other may occur spontaneously. The tendency of such a reaction to occur depends upon the relative affinity of the electron acceptor of each redox pair for electrons. The standard reduction potential, E0, a measure of this affinity, is determined in an experiment such as that described in Figure 13–15. Electrochemists have chosen as a standard of reference the half reaction

H+ + e   →   ½H2

The electrode at which this half-reaction occurs is arbitrarily assigned a standard reduction potential of 0.00 V. When this hydrogen electrode is connected through an external circuit to another half-cell in which the oxidized and reduced species are both present at standard concentrations (each solute at 1 M, each gas at 1 atm), electrons will tend to flow through the external circuit from the half-cell of lower standard reduction potential to the half cell of higher standard reduction potential. By convention, the half-cell with the stronger tendency to acquire electrons is assigned a positive value of E0 (in volts).

The reduction potential of a half-cell depends not only upon the chemical species present but also upon their activities, approximated by their concentrations. About a century ago, Walther Nernst derived an equation that relates standard reduction potential (E0) to reduction

potential (E) at any concentration of oxidized and reduced species in the cell:

where R and T have their usual meanings (Table 13–1), n is the number of electrons transferred per molecule, and ℱ is the Faraday constant, 96.48 kJ/V•mol. At 298 K (25 °C), this expression reduces to:

Many half reactions of interest to biochemists involve protons. As in the definition of ΔG°’, biochemists define the standard state for oxidation–reduction reactions as pH 7 and express reduction potential as E0′, the standard reduction potential at pH 7. The values for standard reduction potentials given in Table 13–7 and used throughout this book are for E0′ and are therefore only valid for calculations involving systems at neutral pH. Each value represents the potential difference when the conjugate redox pair at 1 M concentrations at pH 7 is connected with the standard (pH 0) hydrogen electrode. Notice in Table 13–7 that when the conjugate pair 2H+/H2 at pH 7 is connected with the standard hydrogen electrode (pH 0), electrons tend to flow from the pH 7 cell to the standard (pH 0) cell; the measured ΔE0′ for the 2H+/H2 pair is –0.414 V.

The usefulness of reduction potentials stems from the fact that when E has been determined for any two half-cells, relative to the standard hydrogen electrode, their reduction potentials relative to each other are also known. One can therefore predict the direction in which electrons will tend to flow when these two half-cells are connected through an external circuit, or when the components of the two half-cells are present together in the same solution. Electrons will tend to flow to the half-cell with the more positive E, and the strength of that tendency is proportional to the difference in reduction potentials, ΔE.

The energy made available to do work by this spontaneous electron flow (the free-energy change for the oxidation–reduction reaction) is proportional to ΔE:

ΔG = –n ℱ ΔE,   or   ΔG°’ = –n ℱ ΔE0

Here n represents the number of electrons transferred in the reaction. With this equation it is possible to calculate the free-energy change for any oxidation–reduction reaction from the values of E0′ (found in a table of reduction potentials) and the concentrations of the species involved in the reaction.

Consider the reaction in which acetaldehyde is reduced by the biological electron carrier NADH:

Acetaldehyde + NADH + H+   →   ethanol + NAD+

The relevant half-reactions and their E0′ values (Table 13–7) are:

(1)   Acetaldehyde + 2H+ + 2e   →   ethanol      E0′ = –0.197 V
(2)   NAD+ + 2H+ + 2e   →   NADH + H+ E0′ = –0.320 V

For the overall reaction, ΔE0′ = –0.197 V – (–0.320 V) = 0.123 V, and n is 2. Therefore, ΔG°’ = –n ℱ ΔE0′ = –2 (96.5 kJ/V•mol) (0.123 V) = –23.7 kJ/mol. This is the free-energy change for the oxidation–reduction reaction when acetaldehyde, ethanol, NAD+, and NADH are all present at 1 M concentrations. If, instead, acetaldehyde and NADH were present at 1 M, but ethanol and NAD+ were present at 0.1 M, the value for ΔG would be calculated as follows. First, the values of E for both reductants are determined (Eqn 13–7):

Then ΔE is used to calculate ΔG (Eqn 13–8):

ΔE  = –0.167 V – (–0.350) V = 0.183 V
ΔG  = –n ℱ ΔE
= –2 (96.5 kJ/V•mol) (0.183 V)
= –35.3 kJ/mol

it is thus possible to calculate the free-energy change for any biological oxidation at any concentrations of the redox pairs.

In many organisms, the oxidation of glucose supplies energy for the production of ATP. For the complete oxidation of glucose:

C6H12O6 + 6O2   →   6CO2 + 6H2O

ΔG°’ is –2,840 kJ/mol. This is a much larger change in free energy than that occurring during ATP synthesis (50 to 60 kJ/mol; see Box 13–2). Cells do not convert glucose to CO2 in a single, very energetic reaction, but rather in a series of reactions, some of which are oxidations. The free-energy change of these oxidation steps is larger than, but of the same order of magnitude as, that required for ATP synthesis from ADP. Electrons removed in these oxidation steps are transferred to coenzymes specialized for carrying electrons, such as NAD+ and FAD, which are described below.

Most cells have enzymes to catalyze the oxidation of hundreds of different compounds. These enzymes channel electrons from their substrates into a few types of universal electron carriers. The nucleotides NAD+, NADP+, FMN, and FAD are water-soluble cofactors that undergo reversible oxidation and reduction in many of the electron transfer reactions of metabolism. Their reduction in catabolic processes results in the conservation of free energy released by substrate oxidation. The nucleotides NAD+ and NADP+ move readily from one enzyme to another, but the flavin nucleotides FMN and FAD are very tightly bound to the enzymes, called flavoproteins, for which they serve as prosthetic groups. Lipid-soluble quinones such as ubiquinone and plastoquinone act in the nonaqueous environment of membranes, accepting electrons and conserving free energy. Iron–sulfur proteins and cytochromes are proteins with tightly bound prosthetic groups that undergo reversible oxidation and reduction; they, too, serve as electron carriers in many oxidation–reduction reactions. Some of these proteins are soluble, but others are peripheral or integral membrane proteins (p. 277). We will describe some chemical features of nucleotide cofactors and of certain enzymes (dehydrogenases and flavoproteins) that use them. The oxidation–reduction chemistry of quinones, iron–sulfur proteins, and cytochromes will be discussed in Chapter 18.
Figure 13–16  (a) Nicotinamide adenine dinucleotide (NAD+) and its phosphorylated analog NADP+ undergo reduction to NADH or NADPH, accepting a hydride ion (two electrons and one proton) from an oxidizable substrate. The hydride ion may be added to either the front (A type) or the back (B type) of the planar nicotinamide ring (see Table 13–8). (b) The UV absorption spectra of NAD+ and NADH. Reduction of the nicotinamide ring produces a new, broad absorption band with a maximum at 340 nm. The production of NADH during an enzyme-catalyzed oxidation can be conveniently followed by observing the appearance of the absorbance at 340 nm.

Nicotinamide adenine dinucleotide (NAD+ in its oxidized form) and its close analog nicotinamide adenine dinucleotide phosphate (NADP+) are composed of two nucleotides joined through their phosphate groups by a phosphoric acid anhydride bond (Fig. 13–16). Because their nicotinamide ring resembles pyridine, these compounds are sometimes called pyridine nucleotides. The vitamin niacin provides the nicotinamide moiety for the synthesis of the pyridine nucleotides.

Both coenzymes undergo reversible reduction of the nicotinamide ring (Fig. 13–16). As a substrate molecule undergoes oxidation (dehydrogenation), giving up two hydrogen atoms, the oxidized form of the nucleotide (NAD+ or NADP+) accepts a hydride ion ( ⁚ H, the equivalent

of a proton and two electrons) and is transformed into the reduced form (NADH or NADPH). The second H+ removed from the substrate is released to the aqueous solvent. The half reaction for each nucleotide is therefore

NAD+ + 2e + 2H+   →   NADH + H+
NADP+ + 2e + 2H+   →   NADPH + H+

In the abbreviations NADH and NADPH, the H denotes this added hydride ion, and the loss of the positive charge when H is added to the oxidized form is also made clear. To refer to one of these nucleotides without specifying its oxidation state, we will use NAD or NADP.

The total concentration of NAD+ + NADH in most tissues is about 10–5 M; that of NADP+ + NADP is about 10 times lower. In many cells and tissues, the ratio of NAD+ (oxidized) to NADH (reduced) is high, favoring hydride transfer to NAD+ to form NADH; by contrast, NADPH (reduced) is generally present in greater amounts than its oxidized form, NADP+, favoring hydride transfer from NADPH. This reflects the specialized metabolic roles of the two cofactors: NAD+ generally functions in catabolic oxidations, and NADPH is the usual cofactor in anabolic reductions. A few enzymes will use either cofactor, but most show a strong preference for one cofactor over the other. This functional specialization allows a cell to maintain two distinct pools of electron carriers in the same cellular compartment.

More than 200 enzymes are known to catalyze reactions in which NAD+ or NADP+ accepts a hydride ion from some reduced substrate or NADH or NADPH donates a hydride ion to an oxidized substrate. The general reactions are

AH2 + NAD+   ⇌   A + NADH + H+
AH2 + NADP+   ⇌   A + NADPH + H+

where AH2 is the reduced substrate and A the oxidized substrate. The general name for enzymes of this type is oxidoreductase (see Table 8–3); they are also commonly called dehydrogenases. For example,

the enzyme alcohol dehydrogenase catalyzes the first step in the catabolism of ethanol, in which ethanol is oxidized to acetaldehyde:

CH3CH2OH + NAD+   ⇌   CH3CHO + NADH + H+

Notice that in ethanol, one of the carbon atoms has undergone the loss of hydrogen and has been oxidized from an alcohol to an aldehyde (see Fig. 13–14).

When NAD+ or NADP+ is reduced, the hydride ion could in principle be transferred to either side of the nicotinamide ring: the front (A type) or the back (B type) as represented in Figure 13–16. Studies with isotopically labeled substrates have shown that a given enzyme catalyzes one or the other type of transfer, but not both. For example, yeast alcohol dehydrogenase and lactate dehydrogenase from vertebrate heart both transfer a hydride ion from their respective substrates to the same side of the nicotinamide ring; they are classed as type A dehydrogenases to distinguish them from another group of enzymes that reduce NAD+ by transferring the hydride to the other (B) side of the ring (Fig. 13–16; Table 13–8).

The association between a given dehydrogenase and NAD or NADP is relatively loose; the cofactor readily diffuses from the surface of one enzyme to that of another, acting as a water-soluble carrier of electrons from one metabolite to another. For example, in the production of alcohol during fermentation of glucose by yeast cells, a hydride ion is removed from glyceraldehyde-3-phosphate by one enzyme (glyceraldehyde-3-phosphate dehydrogenase) and transferred to NAD+. The NADH thereby produced then leaves the enzyme surface and diffuses to another enzyme, alcohol dehydrogenase, which transfers a hydride ion from NADH to acetaldehyde, producing ethanol:

(1) Glyceraldehyde-3-phosphate + NAD+   →   3-phosphoglycerate + NADH + H+
(2) Acetaldehyde + NADH + H+   →   ethanol + NAD+

Sum:     Glyceraldehyde-3-phosphate + acetaldehyde   →   3-phosphoglycerate + ethanol

Notice that in the overall reaction there is no net production or consumption of NAD+ or NADH; the cofactors function catalytically, being recycled repeatedly without a net change in the concentration of NAD+ + NADH.

Flavoproteins (Table 13–9) are enzymes that catalyze oxidation–reduction reactions using either flavin mononucleotide (FMN) or flavin adenine dinucleotide (FAD) as cofactor (Fig. 13–17). These cofactors are derived from the vitamin riboflavin. The fused ring structure of flavin nucleotides (the isoalloxazine ring) undergoes reversible reduction, accepting either one or two electrons in the form of hydrogen atoms (electron plus proton) from a reduced substrate; the reduced forms are abbreviated FADH2 and FMNH2. When a fully oxidized flavin nucleotide accepts only one electron (one hydrogen atom), the semiquinone form of the isoalloxazine ring (Fig. 13–17) is produced. Because flavoproteins can participate in either one- or two-electron transfers, this class of proteins is involved in a greater diversity of reactions than the NAD-linked dehydrogenases. As with nicotinamide coenzymes (Fig. 13–16), the reduction of flavin nucleotides is accompanied by a change in a major absorption band. This change can often be used in assaying a reaction involving a flavoprotein.

The flavin nucleotide in most flavoproteins is bound rather tightly and, in some enzymes such as succinate dehydrogenase, covalently. Such tightly bound cofactors are properly called prosthetic groups. They do not carry electrons by diffusing away from one enzyme and to the next; rather, they provide a means by which the flavoprotein can temporarily hold electrons while it catalyzes electron transfer from a reduced substrate to an electron acceptor. One important feature of the flavoproteins is the variability in standard reduction potential (E0′) of the bound flavin nucleotide; tight association between the enzyme and prosthetic group confers on the flavin ring a reduction potential typical of the specific flavoprotein, sometimes quite different from that of the

free flavin nucleotide. Flavoproteins are often very complex; some have, in addition to a flavin nucleotide, tightly bound inorganic ions (iron or molybdenum, for example) capable of participating in electron transfers.

Living cells constantly perform work and thus require energy for the maintenance of highly organized structures, for the synthesis of cellular components, for movement, for the generation of electrical currents, for the production of light, and for many other processes. Bioenergetics is the quantitative study of energy relationships and energy conversions in biological systems. Biological energy transformations obey the laws of thermodynamics. All chemical reactions are influenced by two forces: the tendency to achieve the most stable bonding state (for which enthalpy, H, is a useful expression) and the tendency to achieve the highest degree of randomness, expressed as entropy, S. The net driving force in a reaction is ΔG, the free-energy change, which represents the net effect of these two factors: ΔG = ΔHT ΔS. Cells require sources of free energy to perform work.

The standard free-energy change, ΔG°’, is a physical constant characteristic for a given reaction, and can be calculated from the equilibrium constant for the reaction: ΔG°’ = –RT ln Keq. The actual free-energy change, ΔG, is a variable, which depends on ΔG°’ and on the concentrations of reactants and products: ΔG = ΔG°’ + RT ln ([products]/[reactants]). When ΔG is large and negative, the reaction tends to go in the forward direction; when it is large and positive, the reaction tends to go in the reverse direction; and when ΔG = 0, the system is at equilibrium. The free-energy change for a reaction is independent of the pathway by which the reaction occurs. Free-energy changes are also additive; the net chemical reaction that results from the successive occurrence of reactions sharing a common intermediate has an overall free-energy change that is the sum of the ΔG values for the individual reactions.

ATP is the chemical link between catabolism and anabolism. Its exergonic conversion to ADP and Pi, or to AMP and PPi, is coupled to a large number of endergonic reactions and processes. In general, it is not ATP hydrolysis, but the transfer of phosphate or adenylate from ATP to a substrate or enzyme molecule that couples the energy of ATP breakdown to endergonic transformations of substrates. By these group transfer reactions ATP provides the energy for anabolic reactions, including the synthesis of informational molecules, and for the transport of molecules and ions across membranes against concentration and electrical potential gradients. Muscle contraction is one of several exceptions to this generalization; ATP hydrolysis drives the conformational changes in myosin that produce contraction in muscle.

Cells contain other metabolites with large, negative, free energies of hydrolysis, including phosphoenolpyruvate, 1,3-bisphosphoglycerate, and phosphocreatine. These high-energy compounds, like ATP, have a high phosphate group transfer potential; they are good donors of the phosphate group. Thioesters also have high free energies of hydrolysis.

Biological oxidation–reduction reactions can be described in terms of two half-reactions, each with a characteristic standard reduction potential, E0′. When two electrochemical half-cells, each containing the components of a half-reaction, are connected, electrons tend to flow to the half-cell with the higher reduction potential. The strength of this tendency is proportional to the difference between the two reduction potentials (ΔE), and is a function of the concentrations of oxidized and reduced species. The standard free-energy change for an oxidation–reduction reaction is directly proportional to the difference in standard reduction potentials of the two half cells: ΔG°’ = –n ℱ ΔE0′.

Many biological oxidation reactions are dehydrogenations in which one or two hydrogen atoms (electron and proton) are transferred from a substrate to a hydrogen acceptor. Oxidation–reduction reactions in cells involve specialized electron carrier cofactors. NAD and NADP are the freely diffusible cofactors of many dehydrogenases of cells. Both cofactors accept two electrons and one proton. FAD and FMN, the flavin nucleotides, serve as tightly bound prosthetic groups of flavoproteins. They can accept either one or two electrons. In many organisms, a central energy-conserving process is the stepwise oxidation of glucose to CO2, in which the energy of oxidation is conserved in ATP as electrons are passed to O2.

Further Reading

Bioenergetics and Thermodynamics

Atkins, P.W. (1984) The Second Law, Scientific American Books, Inc., New York. 
A well-illustrated and elementary discussion of the second law and its implications.

Blum, H.F. (1968) Time’s Arrow and Evolution, 3rd edn, Princeton University Press, Princeton, NJ. 

Cantor, C.R. & Schimmel, P.R. (1980) Biophysical Chemistry, W.H. Freeman and Company, San Francisco. 
This and the next two books are outstanding advanced treatments of thermodynamics.

Dickerson, R.E. (1969) Molecular Thermodynamics, W.A. Benjamin, Inc., Menlo Park, CA. 

Edsall, J.T. & Gutfreund, H. (1983) Biothermodynamics: The Study of Biochemical Processes at Equilibrium, John Wiley & Sons, Inc., New York. 

Ingraham, L.L. & Pardee, A.B. (1967) Free energy and entropy in metabolism. In Metabolic Pathways, 3rd edn, Vol. I (Greenberg, D.M., ed), pp. 1–46, Academic Press, Inc., New York. 

Klotz, I.M. (1967) Energy Changes in Biochemical Reactions, Academic Press, Inc., New York. 
Brief and nonmathematical introduction to thermodynamics for biochemists, with many illustrative examples.

Morowitz, H.J. (1970) Entropy for Biologists: An Introduction to Thermodynamics, Academic Press, Inc., New York. 
A good introduction to thermodynamics in biology, not limited to a discussion of entropy.

Rothman, T. (1989) Science à la Mode, Princeton University Press, Princeton, NJ. 
Chapter 4, “The Evolution of Entropy”, is an excellent discussion of entropy in biology.

van Holde, K.E. (1985) Physical Biochemistry, 2nd edn, Prentice-Hall, Inc., Englewood Cliffs, NJ. 
Chapters 1 through 3 cover the thermodynamic concepts discussed in this chapter.

Phosphate Group Transfers and ATP

Alberty, R.A. (1969) Standard Gibbs free energy, enthalpy and entropy changes as a function of pH and pMg for several reactions involving adenosine phosphates. J. Biol. Chem. 244, 3290–3302. 
This research paper documents the strong dependence of the free energy of ATP hydrolysis on the concentrations of H+ and Mg2+.

Bock, R.M. (1960) Adenine nucleotides and properties of pyrophosphate compounds. In The Enzymes, 2nd edn, Vol. 2 (Boyer, P.D., Lardy, H., & Myrback, K., eds), pp. 3–38, Academic Press, Inc., New York. 

Bridger, W.A. & Henderson, J.F. (1983) Cell ATP, John Wiley & Sons, Inc., New York. 
The chemistry of ATP, the role of ATP in metabolic regulation, and the catabolic and anabolic roles of ATP.

Hanson, R.W. (1989) The role of ATP in metabolism. Biochem. Educ. 17, 86–92. 
Excellent summary of the chemistry and biology of ATP.

Harold, F.M. (1986) The Vital Force: A Study of Bioenergetics, W.H. Freeman and Company, New York. 
A beautifully clear discussion of thermodynamics in biological processes.

Jencks, W.P. (1990) How does ATP make work? Chemtracts–Biochem. Mol. Biol. 1, 1–13. 
A clear and sophisticated description of ATP energy transductions in ion transport, muscle contraction, oxidative phosphorylation, and photophosphorylation.

Kalckar, H.M. (1969) Biological Phosphorylations: Development of Concepts, Prentice-Hall, Inc., Englewood Cliffs, NJ. 
An historical account by one of the central participants in the study of biological phosphorylations.

Lipmann, F. (1941) Metabolic generation and utilization of phosphate bond energy. Adv. Enzymol. 11, 99–162. 
The classic description of the role of high-energy phosphate compounds in biology.

Pullman, B. & Pullman, A. (1960) Electronic structure of energy-rich phosphates. Radiat. Res. Suppl. 2, pp. 160–181. 
An advanced discussion of the chemistry of ATP and other “energy-rich” compounds.

Westheimer, F.H. (1987) Why nature chose phosphates. Science 235, 1173–1178. 
A chemist's description of the unique suitability of phosphate esters and anhydrides for metabolic transformations.

Biological Oxidation–Reduction Reactions

Dolphin, D., Avramovic, O., & Poulson, R. (eds) (1987) Pyridine Nucleotide Coenzymes: Chemical, Biochemical, and Medical Aspects, John Wiley & Sons, Inc., New York. 
An excellent two-volume collection of authoritative reviews. Among the most useful of these are the chapters by Kaplan, Westheimer, Veech, and Ohno and Ushio.

Latimer, W.M. (1952) Oxidation Potentials, 2nd edn, Prentice-Hall, Inc., New York. 

Montgomery, R. & Swenson, C.A. (1976) Quantitative Problems in the Biochemical Sciences, 2nd edn, W.H. Freeman and Company, San Francisco. 

Segel, I.H. (1976) Biochemical Calculations, 2nd edn, John Wiley & Sons, Inc., New York. 


1. Entropy Changes during Egg Development  Consider an ecosystem consisting of an egg in an incubator. The white and yolk of the egg contain proteins, carbohydrates, and lipids. If fertilized, the egg is transformed from a single cell to a complex organism. Discuss this irreversible process in terms of the entropy changes in the system, surroundings, and universe. Be sure that you first clearly define the system and surroundings.

2. Calculation of ΔG°’ from Equilibrium Constants  Calculate the standard free-energy changes of the following metabolically important enzyme-catalyzed reactions at 25 °C and pH 7.0 from the equilibrium constants given.

       (a) Glutamate + oxaloacetate     aspartate + α-ketoglutarate     Keq = 6.8
triose phosphate
       (b) Dihydroxyacetone phosphate     glyceraldehyde-3-phosphate     Keq = 0.0475
       (c) Fructose-6-phosphate + ATP     fructose-1,6-bisphosphate + ADP     Keq = 254

3. Calculation of Equilibrium Constants from ΔG°’  Calculate the equilibrium constants Keq for each of the following reactions at pH 7.0 and 25 °C, using the ΔG°’ values of Table 13–4:

       (a) Glucose-6-phosphate + H2O     glucose + Pi
       (b) Lactose + H2O     glucose + galactose
       (c) Malate     fumarate + H2O

4. Experimental Determination of K′eq and ΔG°’  If a 0.1 M solution of glucose-1-phosphate is incubated with a catalytic amount of phosphoglucomutase, the glucose-1-phosphate is transformed to glucose-6-phosphate until equilibrium is established. The equilibrium concentrations are

Glucose-1-phosphate     glucose-6-phosphate
4.5 × 10−3 M 9.6 × 10−2 M

Calculate Keq and ΔG°’ for this reaction at 25 °C.

5. Experimental Determination of ΔG°’ for ATP Hydrolysis  A direct measurement of the standard free-energy change associated with the hydrolysis of ATP is technically demanding because the minute amount of ATP remaining at equilibrium is difficult to measure accurately. The value of ΔG°’ can be calculated indirectly, however, from the equilibrium constants of two other enzymatic reactions having less favorable equilibrium constants:

Glucose-6-phosphate + H2O   →   glucose + Pi       Keq = 270

ATP + glucose   →   ADP + glucose-6-phosphate       Keq = 890

Using this information, calculate the standard free energy of hydrolysis of ATP. Assume a temperature of 25 °C.

6. Difference between ΔG°’ and ΔG  Consider the following interconversion, which occurs in glycolysis (Chapter 14):

Fructose-6-phosphate   ⇌   glucose-6-phosphate       Keq = 1.97

       (a) What is ΔG°’ for the reaction (assuming that the temperature is 25 °C)?

       (b) If the concentration of fructose-6-phosphate is adjusted to 1.5 M and that of glucose-6-phosphate is adjusted to 0.5 M, what is ΔG?
       (c) Why are ΔG°’ and ΔG different?

7. Dependence of ΔG on pH  The free energy released by the hydrolysis of ATP under standard conditions at pH 7.0 is –30.5 kJ/mol. If ATP is hydrolyzed under standard conditions but at pH 5.0, is more or less free energy released? Why?

8. The ΔG°’ for Coupled Reactions  Glucose-1-phosphate is converted into fructose-6-phosphate in two successive reactions:

Glucose-1-phosphate   →   glucose-6-phosphate
Glucose-6-phosphate   →   fructose-6-phosphate

Using the ΔG°’ values in Table 13–4, calculate the equilibrium constant, Keq, for the sum of the two reactions at 25 °C:

Glucose-1-phosphate   →   fructose-6-phosphate

9. Strategy for Overcoming an Unfavorable Reaction: ATP-Dependent Chemical Coupling  The phosphorylation of glucose to glucose-6-phosphate is the initial step in the catabolism of glucose. The direct phosphorylation of glucose by Pi is described by the equation

Glucose + Pi   →   glucose-6-phosphate + H2O       ΔG°’ = 13.8 kJ/mol

       (a) Calculate the equilibrium constant for the above reaction. In the rat hepatocyte the physiological concentrations of glucose and Pi are maintained at approximately 4.8 mM. What is the equilibrium concentration of glucose-6-phosphate obtained by the direct phosphorylation of glucose by Pi? Does this route represent a reasonable metabolic route for the catabolism of glucose? Explain.

       (b) In principle, at least, one way to increase the concentration of glucose-6-phosphate is to drive the equilibrium reaction to the right by increasing the intracellular concentrations of glucose and Pi. Assuming a fixed concentration of Pi at 4.8 mM, how high would the intracellular concentration of glucose have to be to have an equilibrium concentration of glucose-6-phosphate of 250 μM (normal physiological concentration)? Would this route be a physiologically reasonable approach, given that the maximum solubility of glucose is less than 1 M?
       (c) The phosphorylation of glucose in the cell is coupled to the hydrolysis of ATP; that is, part of the free energy of ATP hydrolysis is utilized to effect the endergonic phosphorylation of glucose:

(1) Glucose + Pi   →   glucose-6-phosphate + H2O ΔG°’ = 13.8 kJ/mol
(2) ATP + H2O   →   ADP + Pi   ΔG°’ = –30.5 kJ/mol

Sum:     Glucose + ATP   →   glucose-6-phosphate + ADP

Calculate Keq for the overall reaction. When the ATP-dependent phosphorylation of glucose is carried out, what concentration of glucose is needed to achieve a 250 μM intracellular concentration of glucose-6-phosphate when the concentrations of ATP and ADP are 3.38 and 1.32 mM, respectively? Does this coupling process provide a feasible route, at least in principle, for the phosphorylation of glucose as it occurs in the cell? Explain.
       (d) Although coupling ATP hydrolysis to glucose phosphorylation makes thermodynamic sense, how this coupling is to take place has not been specified. Given that coupling requires a common intermediate, one conceivable route is to use ATP hydrolysis to raise the intracellular concentration of Pi and thus drive the unfavorable phosphorylation of glucose by Pi. Is this a reasonable route? Explain.
       (e) The ATP-coupled phosphorylation of glucose is catalyzed in the hepatocyte by the enzyme glucokinase. This enzyme binds ATP and glucose to form a glucose–ATP–enzyme complex, and the phosphate is transferred directly from ATP to glucose. Explain the advantages of this route.

10. Calculations of ΔG°’ for ATP-Coupled Reactions  From data in Table 13–6 calculate the ΔG°’ value for the reactions

       (a)  Phosphocreatine + ADP   →   creatine + ATP

       (b)  ATP + fructose   →   ADP + fructose-6-phosphate

11. Coupling ATP Cleavage to an Unfavorable Reaction  This problem explores the consequences of coupling ATP hydrolysis under physiological conditions to a thermodynamically unfavorable biochemical reaction. Because we want to explore these consequences in stages, we shall consider the hypothetical transformation, X   →   Y, a reaction for which ΔG°’ = 20 kJ/mol.

       (a) What is the ratio [Y]/[X] at equilibrium?
       (b) Suppose X and Y participate in a sequence of reactions during which ATP is hydrolyzed to ADP and Pi. The overall reaction is:

X + ATP + H2O   →   Y + ADP + Pi

Calculate [Y]/[X] for this reaction at equilibrium. Assume for the purposes of this calculation that the concentrations of ATP, ADP, and Pi are all 1 M when the reaction is at equilibrium.

       (c) We know that [ATP], [ADP], and [Pi] are notM under physiological conditions. Calculate the ratio [Y]/[X] for the ATP-coupled reaction when the values of [ATP], [ADP], and [Pi] are those found in rat myocytes (Table 13–5).

12. Calculations of ΔG at Physiological Concentrations  Calculate the physiological ΔG (not ΔG°’) for the reaction

Phosphocreatine + ADP   →   creatine + ATP

at 25 °C as it occurs in the cytosol of neurons, in which phosphocreatine is present at 4.7 mM, creatine at 1.0 mM, ADP at 0.20 mM, and ATP at 2.6 mM.

13. Free Energy Required for ATP Synthesis under Physiological Conditions  In the cytosol of rat hepatocytes, the mass-action ratio is


  =  5.33 × 102 M–1

Calculate the free energy required to synthesize ATP in the rat hepatocyte.

14. Daily ATP Utilization by Human Adults 

       (a) A total of 30.5 kJ/mol of free energy is needed to synthesize ATP from ADP and Pi when the reactants and products are at 1 M concentration (standard state). Because the actual physiological concentrations of ATP, ADP, and Pi are not 1 M, the free energy required to synthesize ATP under physiological conditions is different from ΔG°’. Calculate the free energy required to synthesize ATP in the human hepatocyte when the physiological concentrations of ATP, ADP, and Pi are 3.5, 1.50, and 5.0 mM, respectively.
       (b) A normal 68 kg (150 lb) adult requires a caloric intake of 2,000 kcal (8,360 kJ) of food per day (24 h). This food is metabolized and the free energy used to synthesize ATP, which is then utilized to do the body’s daily chemical and mechanical work. Assuming that the efficiency of converting food energy into ATP is 50%, calculate the weight of ATP utilized by a human adult in a 24 h period. What percentage of the body weight does this represent?
       (c) Although adults synthesize large amounts of ATP daily, their body weight, structure, and composition do not change significantly during this period. Explain this apparent contradiction.

15. ATP Reserve in Muscle Tissue  The ATP concentration in muscle tissue (approximately 70% water) is about 8.0 mM. During strenuous activity each gram of muscle tissue uses ATP at the rate of 300 μmol/min for contraction.

       (a) How long would the reserve of ATP last during a 100 meter dash?
       (b) The phosphocreatine level in muscle is about 40.0 mM. How does this help extend the reserve of muscle ATP?
       (c) Given the size of the reserve ATP pool, how can a person run a marathon?

16. Rates of Turnover of γ- and β-Phosphates of ATP  If a small amount of ATP labeled with radioactive phosphorus in the terminal position, [γ-32P]ATP, is added to a yeast extract, about half of the 32P activity is found in Pi within a few minutes, but the concentration of ATP remains unchanged. Explain. If the same experiment is carried out using ATP labeled with 32P in the central position, [β-]ATP, the 32P does not appear in Pi within the same number of minutes. Why?

17. Cleavage of ATP to AMP and PPi during Metabolism  The synthesis of the activated form of acetate (acetyl-CoA) is carried out in an ATP-dependent process:

Acetate + CoA + ATP   →   acetyl-CoA + AMP + PPi

       (a) The ΔG°’ for the hydrolysis of acetyl-CoA to acetate and CoA is –32.2 kJ/mol and that for hydrolysis of ATP to AMP and PPi is –30.5 kJ/mol. Calculate ΔG°’ for the ATP-dependent synthesis of acetyl-CoA.

       (b) Almost all cells contain the enzyme inorganic pyrophosphatase, which catalyzes the hydrolysis of PPi to Pi. What effect does the presence of this enzyme have on the synthesis of acetyl-CoA? Explain.

18. Are All Metabolic Reactions at Equilibrium? 

       (a) Phosphoenolpyruvate is one of the two phosphate donors in the synthesis of ATP during glycolysis. In human erythrocytes, the steady-state concentration of ATP is 2.24 mM, that of ADP is 0.25 mM, and that of pyruvate is 0.051 mM. Calculate the concentration of phosphoenolpyruvate at 25 °C, assuming that the pyruvate kinase reaction (Fig. 13–3) is at equilibrium in the cell.
       (b) The physiological concentration of phosphoenolpyruvate in human erythrocytes is 0.023 mM. Compare this with the value obtained in (a). What is the significance of this difference? Explain.

19. Standard Reduction Potentials  The standard reduction potential, E0′, of any redox pair is defined for the half cell reaction:

Oxidizing agent  +  n electrons   ⇌   reducing agent

The E0′ values for the NAD+/NADH and pyruvate/lactate conjugate redox pairs are –0.32 and –0.19 V, respectively.

       (a) Which conjugate pair has the greater tendency to lose electrons? Explain.
       (b) Which is the stronger oxidizing agent? Explain.
       (c) If we begin with 1 M concentrations of each reactant and product at pH 7, in which direction will the following reaction proceed?

Pyruvate + NADH + H+   ⇌   lactate + NAD+

       (d) What is the standard free-energy change (ΔG°’) at 25 °C for this reaction?

       (e) What is the equilibrium constant (Keq) for this reaction?

20. Energy Span of the Respiratory Chain  Electron transfer in the mitochondrial respiratory chain may be represented by the net reaction equation

NADH + H+ + ½O2   ⇌   H2O + NAD+
       (a) Calculate the value of ΔE0′ for the net reaction of mitochondrial electron transfer.
       (b) Calculate ΔG°’ for this reaction.
       (c) How many ATP molecules can theoretically be generated by this reaction if the standard free energy of ATP synthesis is 30.5 kJ/mol?

21. Dependence of Electromotive Force on Concentrations  Calculate the electromotive force (in volts) registered by an electrode immersed in a solution containing the following mixtures of NAD+ and NADH at pH 7.0 and 25 °C, with reference to a half-cell of E0′ = 0.00 V.

       (a) 1.0 mM NAD+ and 10 mM NADH

       (b) 1.0 mM NAD+ and 1.0 mM NADH
       (c) 10 mM NAD+ and 1.0 mM NADH

22. Electron Affinity of Compounds  List the following substances in order of increasing tendency to accept electrons: (a) α-ketoglutarate + CO2 (yielding isocitrate), (b) oxaloacetate, (c) O2, (d) NADP+.

23. Direction of Oxidation–Reduction Reactions  Which of the following reactions would be expected to proceed in the direction shown under standard conditions, assuming that the appropriate enzymes are present to catalyze them?

       (a) Malate + NAD+   →   oxaloacetate + NADH + H+

       (b) Acetoacetate + NADH + H+   →   β-hydroxybutyrate + NAD+

       (c) Pyruvate + NADH + H+   →   lactate + NAD+

       (d) Pyruvate + β-hydroxybutyrate   →   lactate + acetoacetate

       (e) Malate + pyruvate   →   oxaloacetate + lactate

       (f) Acetaldehyde + succinate   →   ethanol + fumarate
Figure 14–1  Major pathways of glucose utilization in cells of higher plants and animals. Although not the only possible fates for glucose, these three pathways are the most significant in terms of the amount of glucose that flows through them in most cells.
Chapter 14
Glycolysis and the Catabolism of Hexoses

Having examined the organizing principles of cell metabolism and bioenergetics, we are ready to see how the chemical energy stored in glucose and other fuel molecules is released to perform biological work. D-Glucose is the major fuel of most organisms and occupies a central position in metabolism. It is relatively rich in potential energy; its complete oxidation to carbon dioxide and water proceeds with a standard free-energy change of –2,840 kJ/mol. By storing glucose as a high molecular weight polymer, a cell can stockpile large quantities of hexose units while maintaining a relatively low cytosolic osmolarity. When the cell’s energy demands suddenly increase, glucose can be released quickly from these intracellular storage polymers.

Glucose is not only an excellent fuel, it is also a remarkably versatile precursor, capable of supplying a huge array of metabolic intermediates, the necessary starting materials for biosynthetic reactions. E. coli can obtain from glucose the carbon skeletons for every one of the amino acids, nucleotides, coenzymes, fatty acids, and other metabolic intermediates needed for growth. A study of the numerous metabolic fates of glucose would encompass hundreds or thousands of transformations. In the higher plants and animals glucose has three major fates: it may be stored (as a polysaccharide or as sucrose), oxidized to a three-carbon compound (pyruvate) via glycolysis, or oxidized to pentoses via the pentose phosphate (phosphogluconate) pathway (Fig. 14–1).

This chapter begins with a description of the individual reactions that constitute the glycolytic pathway and of the enzymes that catalyze them. We then consider fermentation, the operation of the glycolytic pathway under anaerobic conditions. The sources of glucose units for glycolysis are diverse, and we next describe pathways that bring carbon into glycolysis from hexoses other than glucose and from disaccharides and polysaccharides. Like all metabolic pathways, glycolysis is under tight regulation. We discuss the general principles of metabolic regulation, then illustrate these principles with the glycolytic pathway. The chapter concludes with a brief description of two other catabolic pathways that begin with glucose: one leading to pentoses, the other to glucuronate and ascorbic acid (vitamin C).

In glycolysis (from the Greek glykys, meaning “sweet”, and lysis, meaning “splitting”) a molecule of glucose is degraded in a series of

enzyme-catalyzed reactions to yield two molecules of pyruvate. During the sequential reactions of glycolysis some of the free energy released from glucose is conserved in the form of ATP. Glycolysis was the first metabolic pathway to be elucidated and is probably the best understood. From the discovery by Eduard Buchner (in 1897) of fermentation in broken extracts of yeast cells until the clear recognition by Fritz Lipmann and Herman Kalckar (in 1941) of the metabolic role of highenergy compounds such as ATP in metabolism, the reactions of glycolysis in extracts of yeast and muscle were central to biochemical research. The development of methods of enzyme purification, the discovery and recognition of the importance of cofactors such as NAD, and the discovery of the pivotal role in metabolism of phosphorylated compounds all came out of studies of glycolysis. By now, all of the enzymes of glycolysis have been purified from many organisms and thoroughly studied, and the three-dimensional structures of all of the glycolytic enzymes are known from x-ray crystallographic studies.

Glycolysis is an almost universal central pathway of glucose catabolism. It is the pathway through which the largest flux of carbon occurs in most cells. In certain mammalian tissues and cell types (erythrocytes, renal medulla, brain, and sperm, for example), glucose is the sole or major source of metabolic energy through glycolysis. Some plant tissues that are modified for the storage of starch (such as potato tubers) and some plants adapted to growth in areas regularly inundated by water (watercress, for example) derive most of their energy from glycolysis; many types of anaerobic microorganisms are entirely dependent on glycolysis.

Fermentation is a general term denoting the anaerobic degradation of glucose or other organic nutrients into various products (characteristic for different organisms) to obtain energy in the form of ATP. Because living organisms first arose in an atmosphere lacking oxygen, anaerobic breakdown of glucose is probably the most ancient biological mechanism for obtaining energy from organic fuel molecules. In the course of evolution this reaction sequence has been completely conserved; the glycolytic enzymes of vertebrate animals are closely similar, in amino acid sequence and three-dimensional structure, to their homologs in yeast and spinach. The process of glycolysis differs from one species to another only in the details of its regulation and in the subsequent metabolic fate of the pyruvate formed. The thermodynamic principles and the types of regulatory mechanisms in glycolysis are found in all pathways of cell metabolism. A study of glycolysis can serve as a model of many aspects of the pathways discussed later in this book.

Before examining each step of the pathway in some detail, we will take a look at glycolysis as a whole.

Figure 14–2  The two phases of glycolysis. For each molecule of glucose that passes through the preparatory phase (a), two molecules of glyceraldehyde-3-phosphate are formed; both pass through the payoff phase (b). Pyruvate is the end product of the second phase under aerobic conditions, but under anaerobic conditions pyruvate is reduced to lactate to regenerate NAD+. For each glucose molecule, two ATP are consumed in the preparatory phase and four ATP are produced in the payoff phase, giving a net yield of two molecules of ATP per one of glucose converted to pyruvate. The number beside each reaction step corresponds to its numbered heading in the text discussion. Keep in mind that each phosphate group, represented here as Ⓟ has two negative charges (–PO32−).
Figure 14–3  Three possible catabolic fates of the pyruvate formed in the payoff phase of glycolysis. Pyruvate also serves as a precursor in many anabolic reactions, not shown here.

The breakdown of the six-carbon glucose into two molecules of the three-carbon pyruvate occurs in ten steps, the first five of which constitute the preparatory phase (Fig. 14-2a). In these reactions glucose is first phosphorylated at the hydroxyl group on C-6 (step ). The D-glucose-6-phosphate thus formed is converted to D-fructose-6-phosphate (step ), which is again phosphorylated, this time at C-1, to yield D-fructose-1,6-bisphosphate (step ). For both phosphorylations, ATP is the phosphate donor. As all of the sugar derivatives that occur in the glycolytic pathway are the D isomers, we will omit the D designation except when emphasizing stereochemistry.

Fructose-1,6-bisphosphate is next split to yield two three-carbon molecules, dihydroxyacetone phosphate and glyceraldehyde-3-phosphate (step ); this is the “lysis” step that gives the process its name. The dihydroxyacetone phosphate is isomerized to a second molecule of glyceraldehyde-3-phosphate (step ); this ends the first phase of glycolysis. Note that two molecules of ATP must be invested to activate, or prime, the glucose molecule for its cleavage into two three-carbon pieces; later there will be a good return on this investment. To sum up: in the preparatory phase of glycolysis the energy of ATP is invested, raising the free-energy content of the intermediates, and the carbon chains of all the metabolized hexoses are converted into a common product, glyceraldehyde-3-phosphate.

The energetic gain comes in the payoff phase of glycolysis (Fig. 14-2b). Each molecule of glyceraldehyde-3-phosphate is oxidized and phosphorylated by inorganic phosphate (not by ATP) to form 1,3-bisphosphoglycerate (step ). Energy is released as the two molecules of 1,3-bisphosphoglycerate are converted into two molecules of pyruvate (steps through ). Much of this energy is conserved by the coupled phosphorylation of four molecules of ADP to ATP. The net yield is two molecules of ATP per molecule of glucose used, because two molecules of ATP were invested in the preparatory phase of glycolysis. Energy is also conserved in the payoff phase in the formation of two molecules of NADH per molecule of glucose.

In the sequential reactions of glycolysis, three types of chemical transformation are particularly noteworthy: (1) the degradation of the carbon skeleton of glucose to yield pyruvate, (2) the phosphorylation of ADP to ATP by high-energy phosphate compounds formed during glycolysis, and (3) the transfer of hydrogen atoms or electrons to NAD+, forming NADH. The fate of the product, pyruvate, depends on the cell type and the metabolic circumstances.

Fate of Pyruvate  Three alternative catabolic routes are taken by the pyruvate formed by glycolysis. In aerobic organisms or tissues, under aerobic conditions, glycolysis constitutes only the first stage in the complete degradation of glucose (Fig. 14–3). Pyruvate is oxidized, with loss of its carboxyl group as CO2, to yield the acetyl group of acetyl-coenzyme A, which is then oxidized completely to CO2 by the citric acid cycle (Chapter 15). The electrons from these oxidations are passed to O2 through a chain of carriers in the mitochondrion, forming H2O. The energy from the electron transfer reactions drives the synthesis of ATP in the mitochondrion (Chapter 18).

The second route for pyruvate metabolism is its reduction to lactate via lactic acid fermentation. When a tissue such as vigorously contracting skeletal muscle must function anaerobically, the pyruvate cannot be oxidized further for lack of oxygen. Under these conditions pyruvate is reduced to lactate. Certain tissues and cell types (retina, brain, erythrocytes) convert glucose to lactate even under aerobic conditions. Lactate (the dissociated form of lactic acid) is also the product of glycolysis under anaerobic conditions in microorganisms that carry out the lactic acid fermentation (Fig. 14–3).

The third major route for catabolism of pyruvate leads to ethanol. In some plant tissues and in certain invertebrates, protists, and microorganisms such as brewer’s yeast, pyruvate is converted anaerobically into ethanol and CO2, a process called alcohol (or ethanol) fermentation (Fig. 14–3).

The focus of this chapter is catabolism, but pyruvate has other, anabolic, fates. It can, for example, provide the carbon skeleton for the synthesis of the amino acid alanine. We shall return to these anabolic reactions of pyruvate in later chapters.

ATP Formation Coupled to Glycolysis  During glycolysis some of the energy of the glucose molecule is conserved in the form of ATP, while much remains in the product, pyruvate. The overall equation for glycolysis is

Glucose + 2NAD+ + 2ADP + 2Pi   →   2 pyruvate + 2NADH + 2H+ + 2ATP + 2H2O

For each molecule of glucose degraded to pyruvate, two molecules of ATP are generated from ADP and Pi. We can now resolve the equation of glycolysis into two processes: (1) the conversion of glucose into pyruvate, which is exergonic:

Glucose + 2NAD+   →   2 pyruvate + 2NADH + 2H+       ΔG°’1 = –146 kJ/mol

and (2) the formation of ATP from ADP and Pi, which is endergonic:

2ADP + 2Pi   →   2ATP + 2H2O       ΔG°’2 = 2(30.5 kJ/mol) = 61 kJ/mol

If we now write the sum of Equations 14–2 and 14–3, we can also determine the overall standard free-energy change of glycolysis (Eqn 14–1), including ATP formation, as the algebraic sum, ΔG°’s, of ΔG°’1 and ΔG°’2:

ΔG°’s = ΔG°’1 + ΔG°’2 = –146 kJ/mol + 61 kJ/mol = –85 kJ/mol

Under either standard or intracellular conditions, glycolysis is an essentially irreversible process, driven to completion by this large net decrease in free energy. At the actual intracellular concentrations of ATP, ADP, and Pi (see Table 13–5) and of glucose and pyruvate, the efficiency of recovery of the energy of glycolysis in the form of ATP is over 60%.

Energy Remaining in the Pyruvate Produced by Glycolysis  Glycolysis releases only a small fraction of the total available energy of the glucose molecule. When glucose is oxidized completely to CO2 and H2O, the total standard free-energy change is –2,840 kJ/mol. The glycolytic degradation of glucose to two molecules of pyruvate (ΔG°’ =

–146 kJ/mol) therefore yields only (146/2,840)100 = 5.2% of the total energy that can be released by complete oxidation. The two molecules of pyruvate formed by glycolysis still contain most of the biologically available energy of the glucose molecule, energy that can be extracted by oxidative reactions in the citric acid cycle.

Importance of Phosphorylated Intermediates  Each of the nine glycolytic intermediates between glucose and pyruvate is phosphorylated (Fig. 14–2). The phosphate groups appear to have three functions.

       1.  The phosphate groups are ionized at pH 7, thus giving each of the intermediates of glycolysis a net negative charge. Because the plasma membrane is impermeable to molecules that are charged, the phosphorylated intermediates cannot diffuse out of the cell. After the initial phosphorylation, the cell does not have to spend further energy in retaining phosphorylated intermediates despite the large difference between the intracellular and extracellular concentrations of these compounds.

       2.  Phosphate groups are essential components in the enzymatic conservation of metabolic energy. Energy released in the breakage of phosphoric acid anhydride bonds (such as those in ATP) is partially conserved in the formation of phosphate esters such as glucose-6-phosphate. High-energy phosphate compounds formed in glycolysis (1,3-bisphosphoglycerate and phosphoenol pyruvate) donate phosphate groups to ADP to form ATP.

       3.  Binding of phosphate groups to the active sites of enzymes provides binding energy that contributes to lowering the activation energy and increasing the specificity of enzyme-catalyzed reactions (Chapter 8). The phosphate groups of ADP, ATP, and the glycolytic intermediates form complexes with Mg2+, and the substrate binding sites of many of the glycolytic enzymes are specific for these Mg2+ complexes. Nearly all the glycolytic enzymes require Mg2+ for activity.
Figure 14–4  Fate of the carbon atoms of glucose in the formation of glyceraldehyde-3-phosphate.
(a) The origin of the carbons in the two three-carbon products of the aldolase and triose phosphate isomerase reactions. The end product of the two reactions is two molecules of glyceraldehyde-3-phosphate. Each of the three carbon atoms of glyceraldehyde-3-phosphate is derived from either of two specific carbons of glucose (b). The numbering of the carbon atoms of glyceraldehyde-3-phosphate is not identical with the numbering of the carbon atoms of glucose. This is important for interpreting experiments with glucose in which a single carbon is labeled with a radioisotope.

The preparatory phase of glycolysis requires the investment of two molecules of ATP and results in cleavage of the hexose chain into two triose phosphates. The realization that phosphorylated hexoses were intermediates in glycolysis came slowly and serendipitously. In 1906, Arthur Harden and William Young sought to test their hypothesis that inhibitors of proteolytic enzymes would stabilize the glucose-fermenting enzymes in yeast extract. They added blood serum (known to contain inhibitors of proteolytic enzymes) to yeast extracts and observed the predicted stimulation of glucose metabolism. However, in a control experiment intended to show that boiling the serum destroyed the stimulatory activity, they discovered that boiled serum was just as effective at stimulating glycolysis. Careful examination of the contents of the boiled serum revealed that inorganic phosphate was responsible for the stimulation. Harden and Young soon discovered that glucose added to their yeast extract was converted into a hexose bisphosphate (the “Harden-Young ester”, eventually identified as fructose-1,6-bisphosphate). This was the beginning of a long series of investigations of the role of organic esters of phosphate in biochemistry, which has led to our current understanding of the central role of phosphate group transfer in biology.

Phosphorylation of Glucose  In the first step of glycolysis, glucose is primed for subsequent reactions by its phosphorylation at C-6 to yield glucose-6-phosphate; ATP is the phosphate donor:

This reaction, which is irreversible under intracellular conditions, is catalyzed by hexokinase. The common name kinase is applied to enzymes that catalyze the transfer of the terminal phosphate group from ATP to some acceptor – a hexose, in the case of hexokinase. Kinases are a subclass of transferases (see Table 8–3).

Hexokinase catalyzes the phosphorylation not only of D-glucose but also of certain other common hexoses, such as D-fructose and D-mannose. Hexokinase, like many other kinases, requires Mg2+ for its activity, because the true substrate of the enzyme is not ATP4− but the MgATP2− complex (see Fig. 13–2). Detailed studies of the hexokinase of yeast show that the enzyme undergoes a profound change in its shape, an induced fit, when it binds the hexose molecule (see Fig. 8–21). Hexokinase is universally present in cells of all types. Hepatocytes also contain a form of hexokinase called hexokinase D or glucokinase, which is more specific for glucose and differs from other forms of hexokinase in kinetic and regulatory properties (p. 432).

Conversion of Glucose-6-Phosphate to Fructose-6-Phosphate  Phosphohexose isomerase (phosphoglucose isomerase) catalyzes the reversible isomerization of glucose-6-phosphate, an aldose, to yield fructose-6-phosphate, a ketose:

This reaction proceeds readily in either direction, as is predicted from the relatively small change in standard free energy. Phosphohexose isomerase also requires Mg2+ and is specific for glucose-6-phosphate and fructose-6-phosphate.

Phosphorylation of Fructose-6-Phosphate to Fructose-1,6-Bisphosphate  In the second of the two priming reactions of glycolysis, phosphofructokinase-1 catalyzes the transfer of a phosphate group from ATP to fructose-6-phosphate to yield fructose-1,6-bisphosphate:

The reaction is essentially irreversible under cellular conditions. This enzyme is called phosphofructokinase-1 (PFK-1) to distinguish it from a second enzyme (PFK-2; Chapter 19) that catalyzes the formation of fructose-2,6-bisphosphate from fructose-6-phosphate.

In some bacteria and protists, and in most or all plants, there is a phosphofructokinase that uses pyrophosphate (PPi), not ATP, as the phosphate group donor in the synthesis of fructose-1,6-bisphosphate:

Fructose-6-phosphate + PPi     fructose-1,6-bisphosphate + Pi       ΔG°’ = –14 kJ/mol

Phosphofructokinase-1, like hexokinase, is a regulatory enzyme (Chapter 8), one of the most complex known. It is the major point of regulation in glycolysis. The activity of PFK-1 is increased whenever the ATP supply of the cell becomes depleted or when there is an excess of ATP breakdown products, ADP and AMP, particularly the latter. The enzyme is inhibited whenever the cell has ample ATP and when it is well supplied by other fuels such as fatty acids. Fructose-2,6-bisphosphate, structurally similar to the product of this reaction but not an intermediate in glycolysis, is a potent stimulator of both the ATP-dependent and the PPi-dependent enzymes. The regulation of this step in glycolysis is discussed in greater detail later in the chapter.

Cleavage of Fructose-1,6-Bisphosphate  The enzyme fructose-1,6-bisphosphate aldolase, often simply called aldolase, catalyzes a reversible aldol condensation. Fructose-1,6-bisphosphate is cleaved to yield two different triose phosphates, glyceraldehyde-3-phosphate, an aldose, and dihydroxyacetone phosphate, a ketose:

     ΔG°’ = 23.8 kJ/mol
The aldolase of vertebrate animal tissues does not require a divalent cation, but in many microorganisms aldolase is a Zn2+-containing enzyme. Although the aldolase reaction has a strongly positive standard free-energy change in the direction of cleavage, in cells it can proceed readily in either direction. During glycolysis the reaction products (two triose phosphates) are removed quickly by the next two steps, pulling the reaction in the direction of cleavage.

Interconversion of the Triose Phosphates  Only one of the two triose phosphates formed by aldolase – glyceraldehyde-3-phosphate – can be directly degraded in the subsequent reaction steps of glycolysis. However, the other product, dihydroxyacetone phosphate, is rapidly and reversibly converted into glyceraldehyde-3-phosphate by the fifth enzyme of the glycolytic sequence, triose phosphate isomerase:

By this reaction C-1, C-2, and C-3 of the starting glucose now become indistinguishable from C-6, C-5, and C-4, respectively (Fig. 14-4).

This reaction completes the preparatory phase of glycolysis, in which the hexose molecule has been phosphorylated at C-1 and C-6 and then cleaved to form, ultimately, two molecules of glyceraldehyde3-phosphate. Other hexoses, such as D-fructose, D-mannose, and D-galactose, are also convertible into glyceraldehyde-3-phosphate, as we shall see later.
Figure 14–5  (a) A more detailed representation of the glyceraldehyde-3-phosphate dehydrogenase reaction. In step , a covalent thiohemiacetal linkage forms between the substrate and the sulfhydryl group of a Cys residue in the enzyme's active site. This enzyme–substrate intermediate is oxidized by NAD+ (step ), also bound to the active site, converting it into a covalent acyl-enzyme intermediate, a thioester. The enzyme-bound NADH is reoxidized by free NAD+ (step ). The bond between the acyl group and the thiol group of the enzyme has a very high standard free energy of hydrolysis. In step the thioester bond undergoes phosphorolysis (attack by Pi), releasing the free enzyme and an acyl phosphate product (1,3-bisphosphoglycerate), the formation of which conserves much of the free energy liberated during oxidation of the aldehyde. (b) Iodoacetate is a potent inhibitor of glyceraldehyde-3-phosphate dehydrogenase because it forms a covalent derivative of the essential –SH group of the enzyme active site, rendering it inactive.
Figure 14–6  Mechanism of the phosphoglycerate mutase reaction. The enzyme is initially phosphorylated on a His residue by transfer of a phos hate group from 2,3-bisphosphoglycerate. In step of the catalytic reaction, the phosphoenzyme transfers its phosphate group to 3-phosphoglycerate forming 2,3-bisphosphoglycerate. In step the phosphate group at C-3 of 2,3-bisphosphoglycerate is transferred to the same His residue on the enzyme, producing 2-phosphoglycerate and regenerating the phosphoenzyme. The 2,3-bisphosphoglycerate required initially to phosphorylate the enzyme is formed from 3-phosphoglycerate by a specific ATP-dependent kinase; it is then regenerated in step of each catalytic cycle.

The payoff phase of glycolysis (Fig. 14–2b) includes the energy-conserving phosphorylation steps in which some of the free energy of the glucose molecule is conserved in the form of ATP. Remember that one molecule of glucose yields two molecules of glyceraldehyde-3-phosphate; both halves of the glucose molecule follow the same pathway in the second phase of glycolysis. The conversion of two molecules of glyceraldehyde-3-phosphate into two of pyruvate is accompanied by the formation of four molecules of ATP from ADP. However, the net yield of ATP per molecule of glucose degraded is only two, because two molecules of ATP were invested in the preparatory phase of glycolysis to phosphorylate the two ends of the hexose molecule.

Oxidation of Glyceraldehyde-3-Phosphate to 1,3-Bisphosphoglycerate  The first step in the payoff phase of glycolysis is the conversion of glyceraldehyde-3-phosphate to 1,3-bisphosphoglycerate, catalyzed by glyceraldehyde-3-phosphate dehydrogenase:

This is the first of the two energy-conserving reactions of glycolysis that eventually lead to the formation of ATP. The aldehyde group of glyceraldehyde-3-phosphate is dehydrogenated, not to a free carboxyl group, as one might expect, but to a carboxylic acid anhydride with phosphoric acid. This type of anhydride, called an acyl phosphate, has a very high standard free energy of hydrolysis (ΔG°’ = –49.3 kJ/mol; see Fig. 13–4 and Table 13–6). Much of the free energy of oxidation of the aldehyde group of glyceraldehyde-3-phosphate is conserved by formation of the acyl phosphate group at C-1 of 1,3-bisphosphoglycerate.

The acceptor of hydrogen in the glyceraldehyde-3-phosphate dehydrogenase reaction is the coenzyme NAD+ (see Fig. 13–16), the oxidized form of nicotinamide adenine dinucleotide. The reduction of NAD+ proceeds by the enzymatic transfer of a hydride ion ( ⁚ H) from the aldehyde group of glyceraldehyde-3-phosphate to the nicotinamide ring of NAD+, to yield the reduced coenzyme NADH. The other hydrogen atom of the substrate molecule appears in solution as H+ (p. 391).

Oxidation of glyceraldehyde-3-phosphate involves an intermediate in which the substrate is covalently bound to the enzyme (Fig. 14–5a). The aldehyde group of glyceraldehyde-3-phosphate first reacts with the –SH group of an essential Cys residue in the active site of the enzyme. This reaction is homologous with the formation of a hemiacetal (see Fig. 11–6), but in this case the product is a thiohemiacetal. The discovery that glyceraldehyde-3-phosphate dehydrogenase is inhibited by iodoacetate (Fig. 14–5b) was important in the history of research on glycolysis; the addition of this enzyme inhibitor to crude extracts of yeast or muscle caused the accumulation of the hexose phosphates produced in glycolysis, allowing their isolation and identification.

The NADH formed in this step of glycolysis must be reoxidized to NAD+. Cells contain limited amounts of NAD+, and glycolysis would soon come to a halt for lack of NAD+ were the NADH not reoxidized. The reactions in which NAD+ is regenerated anaerobically are described in detail later, in connection with the alternative fates of pyruvate.

Transfer of Phosphate from 1,3-Bisphosphoglycerate to ADP  The enzyme phosphoglycerate kinase transfers the high-energy phosphate group from the carboxyl group of 1,3-bisphosphoglycerate to ADP, forming ATP and 3-phosphoglycerate:

This and the preceding reaction of glycolysis together constitute an energy-coupling process. In these two reactions (steps and ), 1,3-bisphosphoglycerate is the common intermediate; it is formed in the first reaction (which is endergonic), and its acyl phosphate group is transferred to ADP to form ATP in the second reaction (which is strongly exergonic). The sum of these two sequential reactions is

Glyceraldehyde-3-phosphate + ADP + Pi + NAD+   ⇌   3-phosphoglycerate + ATP + NADH + H+
ΔG°’ = –12.5 kJ/mol

Thus the overall reaction is exergonic.

Recall from Chapter 13 that the actual free-energy change, ΔG, is determined by the standard free-energy change, ΔG°’, and the mass-action ratio, which is the ratio [products]/[reactants] (See Eqn 13–3, p. 371). For the first of these two reactions (step )

Notice that [H+] is not included in the mass-action ratio for this reaction. In biochemical calculations [H+] is assumed to be a constant (10–7 M), and this constant is included in the definition of ΔG (Chapter 13).

The second reaction (step ), by consuming the 1,3-bisphosphoglycerate produced in the first, reduces the concentration of 1,3-bisphosphoglycerate and thereby reduces the mass-action ratio for the overall process. When this ratio is less than 1.0, its natural logarithm has a negative sign. If the mass-action ratio is very small, the contribution of the logarithmic term can make ΔG strongly negative. This is simply another way of showing that the two reactions are coupled through a shared intermediate.

The outcome of these two coupled reactions, both reversible under cellular conditions, is that the energy released on oxidation of an aldehyde to a carboxylate group is conserved by the coupled formation of ATP from ADP and Pi. The formation of ATP by phosphate group transfer from a substrate such as 1,3-bisphosphoglycerate is referred to as a substrate-level phosphorylation. We shall later contrast substrate-level phosphorylation with respiration-linked phosphorylation (oxidative phosphorylation), which occurs in mitochondria.

Conversion of 3-Phosphoglycerate to 2-Phosphoglycerate  The enzyme phosphoglycerate mutase catalyzes a reversible shift of the phosphate group between C-2 and C-3 of glycerate. Mg2+ is essential for this reaction:

The reaction occurs in two steps (Fig. 14–6). A phosphate group initially attached to a His residue in the active site of the enzyme is transferred to the hydroxyl group at C-2 of 3-phosphoglycerate, forming 2,3-bisphosphoglycerate. The phosphate at C-3 of 2,3-bisphosphoglycerate is then transferred to the same His residue of the enzyme, producing 2-phosphoglycerate and regenerating the phosphorylated enzyme. Because the enzyme is initially phosphorylated by phosphate transfer from 2,3-bisphosphoglycerate, this compound functions as a cofactor; it is required in small quantities to initiate the catalytic cycle, and is continuously regenerated by that cycle.

Essentially the same mechanism is employed by the enzyme phosphoglucomutase, described below, in the conversion of glucose-1-phosphate into glucose-6-phosphate. In that reaction, glucose-1,6-bisphosphate serves as the essential cofactor. The general name mutase is often given to enzymes that catalyze the transfer of a functional group from one position to another on the same molecule. Mutases are a subclass of isomerases, enzymes that interconvert stereoisomers or structural or positional isomers (see Table 8–3).

Dehydration of 2-Phosphoglycerate to Phosphoenolpyruvate  The second glycolytic reaction that generates a compound with high phosphate group transfer potential is catalyzed by enolase. This enzyme promotes reversible removal of a molecule of water from 2-phosphoglycerate to yield phosphoenolpyruvate:

Despite the relatively small standard free-energy change in this reaction, there is a very large difference in the standard free energy of hydrolysis of the phosphate groups of the reactant and product. That of 2-phosphoglycerate (a low-energy phosphate compound) is –17.6 kJ/mol and that of phosphoenolpyruvate (a super high-energy phosphate compound) is –61.9 kJ/mol (see Fig. 13–3 and Table 13–6). Although 2-phosphoglycerate and phosphoenolpyruvate contain nearly the same total amount of energy, the loss of the water molecule from 2-phosphoglycerate causes a redistribution of energy within the molecule; the standard free-energy change accompanying hydrolysis of the phosphate group is much greater for phosphoenolpyruvate than for 2-phosphoglycerate.

Transfer of the Phosphate Group from Phosphoenolpyruvate to ADP  The last step in glycolysis is the transfer of the phosphate group from phosphoenolpyruvate to ADP, catalyzed by pyruvate kinase:

In this reaction, a substrate-level phosphorylation, the product pyruvate first appears in its enol form. However, the enol form tautomerizes rapidly and nonenzymatically to yield the keto form of pyruvate, the form that predominates at pH 7. The overall reaction has a large, negative standard free-energy change, due in large part to the spontaneous conversion of the enol form of pyruvate into the keto form (see Fig. 13–3). The ΔG°’ of phosphoenolpyruvate hydrolysis is –61.9 kJ/mol; about half of this energy is conserved in the formation of the phosphoric acid anhydride bond of ATP (ΔG°’ = –30.5 kJ/mol) and the rest (–31.4 kJ/mol) constitutes a large driving force pushing the reaction toward ATP synthesis. The pyruvate kinase reaction is essentially irreversible under intracellular conditions. Pyruvate kinase requires K+ and either Mg2+ or Mn2+. It is an important site of regulation, as described later.

We can now construct a balance sheet for glycolysis to account for (1) the fate of the carbon skeleton of glucose, (2) the input of Pi and ADP and the output of ATP, and (3) the pathway of electrons in the oxidation–reduction reactions. The left-hand side of the following equation shows all the inputs of ATP, NAD+, ADP, and Pi (consult Fig. 14–2), and the right-hand side shows all the outputs (keep in mind that each molecule of glucose yields two molecules of glyceraldehyde-3-phosphate):

Glucose + 2ATP + 2NAD+ + 4ADP + 2Pi   →   2 pyruvate + 2ADP + 2NADH + 2H+ + 4ATP + 2H2O

If we cancel out common terms on both sides of the equation, we get the overall equation for glycolysis under aerobic conditions:

Glucose + 2NAD+ + 2ADP + 2Pi   →   2 pyruvate + 2NADH + 2H+ + 2ATP + 2H2O

The two molecules of NADH formed by glycolysis in the cytosol are, under aerobic conditions, reoxidized to NAD+ by transfer of their electrons to the respiratory chain, which in eukaryotic cells is located in the mitochondria. Here these electrons are ultimately passed to O2:

2NADH + 2H+ + O2   →   2NAD+ + 2H2O

Electron transfer from NADH to O2 in mitochondria provides the energy for synthesis of ATP by respiration-linked phosphorylation (Chapter 18).

In the overall process, one molecule of glucose is converted into two molecules of pyruvate (the pathway of carbon). Two molecules of ADP and two of Pi are converted into two molecules of ATP (the pathway of phosphate groups). Four electrons (two hydride ions) are transferred from two molecules of glyceraldehyde-3-phosphate to two of NAD+ (the pathway of electrons).

Figure 14–7  Dilution of a solution containing a noncovalent protein complex favors dissociation of the complex into its constituents.
Figure 14–8  Channeling of a substrate between two enzymes in the glycolytic pathway. When glyceraldehyde-3-phosphate dehydrogenase (blue) and 3-phosphoglycerate kinase (yellow) are combined in vitro, they catalyze the two-step conversion of glyceraldehyde-3-phosphate to 3-phosphoglycerate (Figs. 14–5, 14–6) at a rate greater than the rate at which the first step is catalyzed in the presence of the first enzyme only. Apparently the transfer of 1,3-bisphosphoglycerate from the surface of the dehydrogenase to that of the kinase is faster than the dissociation of 1,3-bisphosphoglycerate from the dehydrogenase into the surrounding medium (which occurs in the absence of the kinase). Physical studies show that the two enzymes can form a stable complex, as is required for substrate channeling between them.
Sequential action of two separate enzymes: the product of the first enzyme (1,3-bisphosphoglycerate) diffuses to the second enzyme. Substrate channeling through a functional complex of two enzymes: the intermediate (1,3-bisphosphoglycerate) is never released to the solvent.

Although the enzymes of glycolysis usually are described as soluble components of the cytosol, there is growing evidence that within the cell these enzymes exist as multienzyme complexes. The classic approach of enzymology – the purification of individual proteins from extracts of broken cells – was applied with great success to the enzymes of glycolysis; we have noted that each of the enzymes has been purified to homogeneity. However, the first casualty of cell breakage is higher-level organization within a cell – the noncovalent and reversible interaction of one protein with another, or of an enzyme with some structural component such as a membrane, microtubule, or microfilament. When cells are broken open, their contents, including enzymes, undergo dilution by a factor of a hundred or a thousand (Fig. 14–7).

When the purified enzymes of glycolysis are combined in vitro at relatively high concentrations, they form specific, functional aggregates, which may reflect their true state inside cells. Several types of evidence suggest that such complexes act in cells to ensure efficient passage of the product of one enzyme to the next enzyme in the pathway for which that product serves as substrate. Kinetic evidence for the channeling of 1,3-bisphosphoglycerate from glyceraldehyde-3-phosphate

dehydrogenase to phosphoglycerate kinase without entering solution (Fig. 14–8) is corroborated by physical evidence that these two enzymes form stable, noncovalent complexes. There is similar evidence for channeling of intermediates between other glycolytic enzymes, such as glyceraldeyde-3-phosphate from aldolase to glyceraldehyde-3-phosphate dehydrogenase.

Furthermore, certain glycolytic enzymes form speciiic noncovalent complexes with structural components of the cell, which may serve to organize reaction sequences and assure efficient transfer of intermediates between cellular compartments. Certain glycolytic enzymes bind to microtubules or to actin microfilaments (see Fig. 2–18), bringing those enzymes into close association and holding them in a specific region of the cytoplasm. Hexokinase binds specifically to the outer membrane of mitochondria. This association may allow ATP produced within the mitochondrion to move directly to the catalytic site of hexokinase without entering, and being diluted by, the cytosol. There is strong evidence for substrate channeling through multienzyme complexes in other metabolic pathways, and it seems likely that many enzymes now thought of as “soluble” actually function in the cell as highly organized complexes that channel intermediates.

During his studies of the fermentation of glucose by yeast, Louis Pasteur discovered that both the rate and the total amount of glucose consumption were many times greater under anaerobic conditions than under aerobic conditions. Later studies of muscle showed the same large difference in the rate of glycolysis under anaerobic and aerobic conditions. The biochemical basis of this “Pasteur effect” is now clear. The ATP yield from glycolysis under anaerobic conditions (2 ATP per molecule of glucose) is much smaller than that from the complete oxidation of glucose to CO2 under aerobic conditions (36 or 38 ATP per glucose molecule; see Chapter 18). About 18 times as much glucose must therefore be consumed anaerobically as aerobically to yield the same amount of ATP.

The flux of glucose through the glycolytic pathway is regulated to achieve constant ATP levels (as well as adequate supplies of glycolytic intermediates that serve biosynthetic roles). The required adjustment in the rate of glycolysis is achieved by the regulation of two glycolytic enzymes: phosphofructokinase-1 and pyruvate kinase. Both enzymes are regulated allosterically by second-to-second fluctuations in the concentration of certain key metabolites that reflect the cellular balance between ATP production and consumption. We return to a more detailed discussion of the regulation of glycolysis later in the chapter.

Pyruvate, the product of glycolysis, represents an important junction point in carbohydrate catabolism (Fig. 14–3). Under aerobic conditions pyruvate is oxidized to acetate, which enters the citric acid cycle (Chapter 15) and is oxidized to CO2 and H2O. The NADH formed by the

dehydrogenation of glyceraldehyde-3-phosphate is reoxidized to NAD+ by passage of its electrons to O2 in the process of mitochondrial respiration (Chapter 18). However, under anaerobic conditions (as in very active skeletal muscles, in submerged plants, or in lactic acid bacteria, for example), NADH generated by glycolysis cannot be reoxidized by O2. Failure to regenerate NAD+ would leave the cell with no electron acceptor for the oxidation of glyceraldehyde-3-phosphate, and the energy-yielding reactions of glycolysis would stop. NAD+ must therefore be regenerated by some other reaction.

The earliest cells to arise during evolution lived in an atmosphere almost devoid of oxygen and had to develop strategies for carrying out glycolysis under anaerobic conditions. Most modern organisms have retained the ability to continually regenerate NAD+ during anaerobic glycolysis by transferring electrons from NADH to form a reduced end product such as lactate or ethanol.

When animal tissues cannot be supplied with sufiicient oxygen to support aerobic oxidation of the pyruvate and NADH produced in glycolysis, NAD+ is regenerated from NADH by the reduction of pyruvate to lactate. Certain other tissues and cell types (retina, brain, erythrocytes) also produce lactate from glucose under aerobic conditions; lactate is a major product of erythrocyte metabolism. The reduction of pyruvate is catalyzed by lactate dehydrogenase, which forms the L isomer of lactic acid (lactate at pH 7). The overall equilibrium of this reaction strongly favors lactate formation, as shown by the large negative standard free-energy change. In glycolysis, dehydrogenation of the two molecules of glyceraldehyde-3-phosphate derived from each molecule of glucose converts two molecules of NAD+ to two of NADH. Because the reduction of two molecules of pyruvate to two of lactate regenerates two molecules of NAD+, the overall process is balanced and can continue indefinitely: one molecule of glucose is converted to two of lactate, with the generation of two ATP molecules from one of glucose, and NAD+ and NADH are continuously interconverted with no net gain or loss in the amount of either.

Although there are two oxidation–reduction steps as glucose is converted into lactate, there is no net change in the oxidation state of carbon; in glucose (C6H12O6) and lactic acid (C3H6O3), the H : C ratio is the same. Nevertheless, some of the energy of the glucose molecule has been extracted by its conversion to lactate, enough to give a net yield of two molecules of ATP for every one of glucose consumed. The lactate formed by active muscles of vertebrate animals can be recycled; it is carried in the blood to the liver where it is converted into glucose during the recovery from strenuous muscle activity (Box 14–1).

Many microorganisms ferment glucose and other hexoses to lactate. Certain lactobacilli and streptococci, for example, ferment the lactose in milk to lactic acid. The dissociation of lactic acid to lactate and H+ in the fermentation mixture lowers the pH, denaturing casein and other milk proteins and causing them to precipitate. Under the correct conditions, the resultant curdling produces cheese or yogurt, depending on which microorganism is involved.

B O X  14–1
Glycolysis without Oxygen: Alligators and Coelacanths

Most vertebrates are essentially aerobic organisms; they first convert glucose into pyruvate by glycolysis and then oxidize the pyruvate completely to CO2 and H2O using molecular oxygen. Anaerobic catabolism of glucose (fermentation to lactate) occurs in most vertebrates, including human beings, during short bursts of extreme muscular activity, for example in a 100 m sprint, during which oxygen cannot be carried to the muscles fast enough to oxidize pyruvate for generating ATP. Instead, the muscles use their stored glycogen as fuel to generate ATP by fermentation, with lactate as the end product. In a sprint, the lactate in the blood builds up to high concentrations. It is slowly converted back into glucose by gluconeogenesis in the liver in the subsequent rest or recovery period, during which oxygen is consumed at a gradually diminishing rate until the breathing rate returns to normal. The excess oxygen consumed in the recovery period represents the repayment of the oxygen debt. This is the amount of oxygen required to supply ATP for gluconeogenesis during recovery respiration, in order to regenerate the glycogen “borrowed” from liver and muscle to carry out intense muscular activity in the sprint. The cycle of reactions that includes glucose conversion to lactate in muscle and lactate conversion to glucose in liver is called the Cori cycle, for Carl and Gerty Cori, whose studies in the 1930s and 1940s clarified the pathway and its role.

The circulatory systems of most small vertebrates can carry oxygen to their muscles fast enough to avoid having to use muscle glycogen anaerobically. For example, migrating birds often fly great distances at high speeds without rest and without incurring an oxygen debt. Many running animals of moderate size also have an essentially aerobic metabolism in their skeletal muscle. However in larger animals, including humans, the circulatory system cannot completely sustain aerobic metabolism in skeletal muscles during long bursts of muscular activity. Such animals generally are slow-moving under normal circumstances and engage in intense muscular activity only in the gravest emergencies, because such bursts of activity require long recovery periods to repay the oxygen debt.

Alligators and crocodiles, for example, are normally sluggish and torpid. Yet when provoked these animals are capable of lightning-fast charges and dangerous lashings of their powerful tails. Such intense bursts of activity are short and must be followed by long periods of recovery. The fast emergency movements require lactate fermentation to generate ATP in skeletal muscles. Because the stores of muscle glycogen are not large, they are rapidly expended in intense muscular activity. Moreover, in such bursts of action, lactate reaches very high concentration in muscles and extracellular fluid. Whereas a trained athlete can recover from a 100 m sprint in 30 min or less, an alligator may require many hours of rest and extra oxygen consumption to clear the excess lactate from its blood and regenerate muscle glycogen.

Other large animals, such as the elephant and rhinoceros, have similar metabolic problems, as do diving mammals such as whales and seals. Dinosaurs and other huge, now-extinct animals probably had to depend on lactate fermentation to supply energy for muscular activity, followed by very long recovery periods during which they were vulnerable to attack by smaller predators better able to use oxygen and thus better adapted to continuous, sustained muscular activity.

Deep-sea explorations have revealed many species of marine life at great ocean depths, where the oxygen concentration is near zero. For example, the primitive coelacanth, a large fish recovered from depths of 4,000 m or more off the coast of South Africa, has been found to have an essentially anaerobic metabolism in virtually all its tissues. It converts carbohydrates by anaerobic mechanisms into lactate and other products, most of which must be excreted. Some marine vertebrates ferment glucose to ethanol and CO2 in order to obtain energy in the form of ATP.

Yeast and other microorganisms ferment glucose to ethanol and CO2, rather than to lactate. Glucose is converted to pyruvate by glycolysis, and the pyruvate is converted to ethanol and CO2 in a two-step process. In the first step, pyruvate undergoes decarboxylation in the irreversible reaction catalyzed by pyruvate decarboxylase. This reaction is a simple decarboxylation and does not involve the net oxidation of pyruvate. Pyruvate decarboxylase requires Mg2+ and has a tightly bound coenzyme, thiamine pyrophosphate, discussed in more detail below.

In the second step, acetaldehyde is reduced to ethanol, with NADH derived from glyceraldehyde-3-phosphate dehydrogenation furnishing the reducing power, through the action of alcohol dehydrogenase. Ethanol and CO2, instead of lactate, are thus the end products of alcohol fermentation. The overall equation of alcohol fermentation is

Glucose + 2ADP + 2Pi   →   2 ethanol + 2CO2 + 2ATP + 2H2O

As in lactic acid fermentation, there is no net change in the ratio of hydrogen to carbon atoms when glucose (H : C ratio = 12/6 = 2) is fermented

to two ethanol and two CO2 (combined H : C ratio = 12/6 = 2). In all fermentations, the H : C ratio of the reactants and products remains the same.

Pyruvate decarboxylase is characteristically present in brewer’s and baker’s yeast and in all other organisms that promote alcohol fermentation, including some plants. The CO2 produced by pyruvate decarboxylation in brewer’s yeast is responsible for the characteristic carbonation of champagne. The ancient art of brewing beer involves a number of enzymatic processes in addition to the reactions of alcohol fermentation (Box 14–2). In baking, CO2 released by pyruvate decarboxylase when yeast is mixed with a fermentable sugar causes dough to rise. The enzyme is absent in the tissues of vertebrate animals and in other organisms, such as the lactic acid bacteria, that carry out lactic acid fermentation.

Alcohol dehydrogenase is present in many organisms that metabolize alcohol, including humans. In human liver it brings about the oxidation of ethanol, either ingested or produced by intestinal microorganisms, with the concomitant reduction of NAD+ to NADH.

B O X  14–2
Brewing Beer

Beer is made by alcohol fermentation of the carbohydrates present in cereal grains (seeds) such as barley, but these carbohydrates, largely polysaccharides, are not available to the glycolytic enzymes in yeast cells until they have been degraded to disaccharides and monosaccharides. The barley must first undergo a process called malting. The cereal seeds are allowed to germinate until they form the hydrolytic enzymes required to break down the polysaccharides of their cell walls and the starch and other polysaccharide food reserves. Germination is then stopped by controlled heating, before further growth of the seedlings occurs. The product is malt, which now contains enzymes such as α-amylase and maltase, capable of breaking down starch to maltose, glucose, and other simple sugars. The malt also contains enzymes specific for the β linkages of cellulose and other cell-wall polysaccharides of the barley husks, which must be broken down in order to allow α-amylase to act on the starch within the seeds.

In the next step the brewer prepares the wort, the nutrient medium required for the subsequent fermentation by yeast cells. The malt is mixed with water and then mashed or crushed. This allows the enzymes formed in the malting process to act on the cereal polysaccharides to form maltose, glucose, and other simple sugars, which are soluble in the aqueous medium. The remaining cell matter is then separated, and the liquid wort is boiled with hops, to give flavor. The wort is cooled and then aerated.

Now the yeast cells are added. In the aerobic wort the yeast grows and reproduces very rapidly, using energy obtained from some of the sugars in the wort. In this phase no alcohol is formed because the yeast, being amply supplied with oxygen, oxidizes the pyruvate formed by glycolysis to CO2 and H2O via the citric acid cycle. When all the dissolved oxygen in the vat of wort has been consumed, the yeast cells switch to anaerobic metabolism of the sugar. From this point on, the yeast ferments the sugars of the wort into ethanol and CO2. The fermentation process is controlled in part by the concentration of the ethanol formed, by the pH, and by the amount of remaining sugar. After the fermentation has been stopped, the cells are removed, and the “raw” beer is ready for final processing.

In the final steps of brewing, the amount of foam or head on the beer, which results from dissolved proteins, is adjusted. Normally this is controlled by the action of proteolytic enzymes that appear in the malting process. If these enzymes act on the beer proteins too long, the beer will have very little head and will be flat; if they do not act long enough, the beer will not be clear when it is cold. Sometimes proteolytic enzymes from other sources are added to control the head.

Figure 14–9  (a) Thiamine pyrophosphate (TPP), the coenzyme form of vitamin B1 (thiamin). The reactive carbon atom in the thiazolium ring is shown in red. In the reaction catalyzed by pyruvate decarboxylase, two of the three carbons of pyruvate are carried transiently on TPP in the form of hydroxyethyl thiamine pyrophosphate (b). This “active acetaldehyde” group (in red) is subsequently released as acetaldehyde.
(c) The cleavage of a carbon–carbon bond often leaves behind a free electron pair or carbanion on one of the products. The strong tendency of a carbanion to form a new bond generally renders a carbanion intermediate unstable. The thiazolium ring of TPP stabilizes carbanion intermediates by providing an electrophilic (electron-deficient) structure into which the carbanion electrons can be delocalized by resonance. Structures with this property are often called “electron sinks”, and they play a role in many biochemical reactions. This principle is illustrated here for the reaction catalyzed by pyruvate decarboxylase. In step , the TPP carbanion acts as a nucleophile, adding to the carbonyl group of pyruvate. In step , a carbanion is formed following decarboxylation. The thiazolium ring acts as an electron sink, stabilizing the carbanion by resonance. After protonation (step ), the reaction product acetaldehyde is released (step ).

The pyruvate decarboxylase reaction in alcohol fermentation represents our first encounter with thiamine pyrophosphate (TPP) (Fig. 14–9), a coenzyme derived from vitamin B1. The absence of vitamin B1 in the human diet leads to the condition known as beriberi, characterized by an accumulation of body fluids (swelling), pain, paralysis, and ultimately death.

Thiamine pyrophosphate plays an important role in the cleavage of bonds adjacent to a carbonyl group (such as the decarboxylation of α-keto acids) and in chemical rearrangements involving transfer of an activated aldehyde group from one carbon atom to another (Table 14–1). The functional part of thiamine pyrophosphate is the thiazolium ring (Fig. 14–9a). The proton at C-2 of the ring is relatively acidic, and loss of this acidic proton produces a carbanion that is the active species in TPP-dependent reactions (Fig. 14–9c). This carbanion readily adds to carbonyl groups, and the thiazolium ring is thereby positioned to act as an “electron sink” that greatly facilitates reactions such as the decarboxylation catalyzed by pyruvate decarboxylase.

Figure 14–10  An industrial-scale fermentation. Microorganisms are cultured in a sterilizable vessel containing thousands of liters of growth medium made up of an inexpensive carbon-and-energy source under carefully controlled conditions, including low oxygen concentration and constant temperature. After centrifugal separation of the cells from the growth medium, the valuable products of the fermentation are recovered from the cells or the supernatant fluid.

Although lactate and ethanol are common products of microbial fermentations, they are by no means the only possible ones. In 1910 Chaim Weizmann (later to become the first president of Israel) discovered that a bacterium, Clostridium acetobutyricum, ferments starch to butanol and acetone. This discovery opened the field of industrial fermentations, in which some readily available material rich in carbohydrate (corn starch or molasses, for example) is supplied to a pure culture of a specific microorganism, which ferments it into a product of greater value. The methanol used to make “gasohol” is produced by microbial fermentation, as are formic, acetic, propionic, butyric, and succinic acids, glycerol, isopropanol, butanol, and butanediol. Fermentations such as these are generally carried out in huge, closed vats in which temperature and access to air are adjusted to favor the multiplication

of the desired microorganism and to exclude contaminating organisms (Fig. 14–10). The beauty of industrial fermentations is that complicated, multistep chemical transformations are carried out in high yields and with few side products by chemical factories that reproduce themselves – microbial cells. In some cases it is possible to immobilize the cells in an inert support, to pass the starting material continuously through a bed of immobilized cells, and to collect the desired product in the effluent: an engineer's dream!

In addition to glucose, many other carbohydrates ultimately enter the glycolytic pathway to undergo energy-yielding degradation. The most significant are the storage polysaccharides glycogen and starch, the disaccharides maltose, lactose, trehalose, and sucrose, and the monosaccharides fructose, mannose, and galactose. We shall now consider the pathways by which these carbohydrates can enter glycolysis.

Figure 14–11  Removal of a terminal glucose residue from the nonreducing end of a glycogen chain by the action of glycogen phosphorylase. This process is repetitive, removing successive glucose residues until it reaches the fourth glucose unit from a branch point (see Fig. 14–12). Amylopectin is degraded in a similar fashion by starch phosphorylase.
Figure 14–12  Glycogen breakdown near (α1→6) branch points. Following the sequential removal of terminal glucose residues by glycogen phosphorylase (Fig. 14–11), glucose residues near a branch are removed in a two-step process that requires the action of a bifunctional “debranching enzyme”. First, the transferase activity of this enzyme shifts a block of three glucose residues from the branch to a nearby nonreducing end, to which they are reattached in (α1→4) linkage. Then the single glucose residue remaining at the branch point, in (α1→6) linkage, is released as free glucose by the enzyme’s (α1→6) glucosidase activity. The glucose residues are shown in shorthand form, which omits the –H, –OH, and –CH2OH groups from the pyranose rings.

The glucose units of the outer branches of glycogen and starch gain entrance into the glycolytic pathway through the sequential action of two enzymes: glycogen phosphorylase (or the similar starch phosphorylase in plants) and phosphoglucomutase. Glycogen phosphorylase catalyzes the reaction in which an (α1→4) glycosidic linkage joining two glucose residues in glycogen undergoes attack by inorganic phosphate, removing the terminal glucose residue as α-D-glucose-1-phosphate (Fig. 14–11). This phosphorolysis reaction that occurs during intracellular mobilization of glycogen stores is different from the hydrolysis of glycosidic bonds by amylase during intestinal degradation of glycogen or starch; in phosphorolysis, some of the energy of the glycosidic bond is preserved in the formation of the phosphate ester, glucose-1-phosphate.

Pyridoxal phosphate is an essential cofactor in the glycogen phosphorylase reaction; its phosphate group acts as a general acid catalyst, promoting attack by Pi on the glycosidic bond. A quite different role of pyridoxal phosphate as a cofactor in amino acid metabolism will be described in detail in Chapter 17.

Glycogen phosphorylase (or starch phosphorylase) acts repetitively on the nonreducing ends of glycogen (or amylopectin) branches until it reaches a point four glucose residues away from an (α1→6) branch point (see Fig. 11–15). Here the action of glycogen or starch phosphorylase stops. Further degradation can occur only after the action of a “debranching enzyme”, oligo (α1→6) to (α1→4) glucantransferase, which catalyzes two successive reactions that remove branches (Fig. 14–12).

Glucose-1-phosphate, the end product of the glycogen and starch phosphorylase reactions, is converted into glucose-6-phosphate by phosphoglucomutase, which catalyzes the reversible reaction

Glucose-1-phosphate   ⇌   glucose-6-phosphate

Phosphoglucomutase requires as a cofactor glucose-1,6-bisphosphate; its role is analogous to that of 2,3-bisphosphoglycerate in the reaction catalyzed by phosphoglycerate mutase (Fig. 14-6). Phosphoglucomutase, like phosphoglycerate mutase, cycles between a phosphorylated and nonphosphorylated form. In phosphoglucomutase, however, it is the hydroxyl group of a Ser residue in the active site that is transiently phosphorylated in the catalytic cycle.

Figure 14–13  Pathway of the conversion of ngalactose into D-glucose. The conversion proceeds through a sugar-nucleotide derivative, UDP-galactose, which is formed when galactose-1-phosphate displaces glucose-1-phosphate from UDP-glucose. UDP-galactose is then converted by UDP-glucose 4-epimerase to UDP-glucose. The UDP-glucose is recycled through another round of the same reaction. The net effect of this cycle is the conversion of galactose-1-phosphate to glucose-1-phosphate; there is no net production or consumption of UDP-galactose or UDP-glucose.

In most organisms, hexoses other than glucose can undergo glycolysis after conversion to a phosphorylated derivative. D-Fructose, present in free form in many fruits and formed by hydrolysis of sucrose in the small intestine, can be phosphorylated by hexokinase, which acts on a number of different hexoses:

Fructose + ATP     fructose-6-phosphate + ADP

In the muscles and kidney of vertebrates this is a major pathway. In the liver, however, fructose gains entry into glycolysis by a different pathway. The liver enzyme fructokinase catalyzes the phosphorylation of fructose, not at C-6, but at C-1:

Fructose + ATP     fructose-1-phosphate + ADP

The fructose-1-phosphate is then cleaved to form glyceraldehyde and dihydroxyacetone phosphate by fructose-1-phosphate aldolase.

Dihydroxyacetone phosphate is converted into glyceraldehyde-3-phosphate by the glycolytic enzyme triose phosphate isomerase. Glyceraldehyde is phosphorylated by ATP and triose kinase to glyceraldehyde-3-phosphate:

Glyceraldehyde + ATP     glyceraldehyde-3-phosphate + ADP

Thus both products of fructose hydrolysis enter the glycolytic pathway as glyceraldehyde-3-phosphate.

D-Galactose, derived by hydrolysis of the disaccharide lactose (milk sugar), is first phosphorylated at C-1 at the expense of ATP by the enzyme galactokinase:

Galactose + ATP   →   galactose-1-phosphate + ADP

The galactose-1-phosphate is then converted into its epimer at C-4, glucose-1-phosphate, by a set of reactions in which uridine diphosphate (UDP) functions as a coenzymelike carrier of hexose groups (Fig. 14–13).

There are several human genetic diseases in which galactose metabolism is affected. In the most common form of galactosemia, the enzyme UDP-glucose:galactose-1-phosphate uridylyltransferase (Fig. 14–13) is genetically defective, preventing the overall conversion of galactose into glucose. Other forms of galactosemia result when either galactokinase or UDP-glucose-4-epimerase is genetically defective.

D-Mannose, which arises from the digestion of various polysaccharides and glycoproteins present in foods, can be phosphorylated at C-6 by hexokinase:

Mannose + ATP     mannose-6-phosphate + ADP

Mannose-6-phosphate is then isomerized by the action of phosphomannose isomerase, to yield fructose-6-phosphate, an intermediate of glycolysis.
Figure 14–14  Lactase, a disaccharidase of the intestinal epithelium, can be detected by treating a thin section of intestinal tissue with an antibody that specifically binds to the enzyme. The antibodies are made visible in the electron microscope by attaching to them tiny colloidal particles of gold, which appear as black (electron-dense) dots in electron micrographs. (a) Tissue from an adult who has retained high levels of lactase. Microvilli are heavily labeled with antibodies that detect lactase. (b) Intestinal microvilli in tissue from an adult with lactose intolerance are much less heavily labeled with antibodies against lactase.

Disaccharides cannot directly enter the glycolytic pathway; indeed they cannot enter cells without first being hydrolyzed to monosaccharides extracellularly. In vertebrates, ingested disaccharides must first be hydrolyzed by enzymes attached to the outer surface of the epithelial cells lining the small intestine (Fig. 14–14), to yield their monosaccharide units:

Maltose + H2O     2 D-glucose
Lactose + H2O     D-galactose + D-glucose
Sucrose + H2O     D-fructose + D-glucose
Trehalose + H2O     2 D-glucose

The monosaccharides so formed are transported into the cells lining the intestine, from which they pass into the blood and are carried to the liver. There they are phosphorylated and funneled into the glycolytic sequence as described above.

Lactose intolerance is a condition, common among adults of most human races except Northern Europeans and some Africans, in which the ingestion of milk or other foods containing lactose leads to abdominal cramps and diarrhea. Lactose intolerance is due to the disappearance after childhood of most or all of the lactase activity of the intestinal cells (Fig. 14–14b), so that lactose cannot be completely digested and absorbed. Lactose not absorbed in the small intestine is converted by bacteria in the large intestine into toxic products that cause the symptoms of the condition. In those parts of the world where lactose intolerance is prevalent, milk is simply not used as a food by adults. Milk products digested with lactase are commercially available in some countries as an alternative to excluding milk products from the diet. In certain diseases of humans, several or all of the intestinal disaccharidases are missing because of genetic defects or dietary factors, resulting in digestive disturbances triggered by disaccharides in the diet (Fig. 14–14b). Altering the diet to reduce disaccharide content sometimes alleviates the symptoms of these defects.

Figure 14–15 summarizes the feeder pathways that funnel hexoses, disaccharides, and polysaccharides into the central glycolytic pathway.

Figure 14–15  Entry of glycogen, starch, disaccharides, and hexoses into the preparatory stage of glycolysis.

Carbohydrate catabolism provides ATP as well as precursors for a variety of biosynthetic processess. It is crucial to a cell to maintain a sufficient concentration of ATP, at a nearly constant level, regardless of which fuel is used to produce ATP and regardless of the rate at which ATP is consumed. An organism that undergoes a change in circumstances, such as increased muscular activity, decreased availability of oxygen, or decreased dietary intake of carbohydrate, must alter its catabolic patterns to change the flow of carbohydrate fuel, whether from stored reserves or from extracellular sources, through glycolysis. These changes in catabolic patterns are accomplished by the regulation of key enzymes in the catabolic pathways. In glycolysis in muscle and liver tissue, four enzymes play a regulatory role: glycogen phosphorylase, hexokinase, phosphofructokinase-1, and pyruvate kinase. Our discussion of the regulation of glycolysis necessarily involves some details of the reciprocally regulated process of glucose synthesis (gluconeogenesis), which is more fully discussed in Chapter 19.

Before describing the regulation of glucose catabolism, we will consider some general principles that apply to the regulation of all biochemical pathways.

Table 14–2  Cytosolic concentrations of enzymes and metabolites of
the glycolytic pathway in skeletal muscle
Enzyme                    Concentration
Metabolite                Concentration
Triose phosphate isomerase
Phosphoglycerate kinase
Phosphoglycerate mutase
Pyruvate kinase
Lactate dehydrogenase
Dihydroxyacetone phosphate

Source: From Srivastava, D.K. & Bernhard, S.A. (1987) Biophysical chemistry of metabolic reaction
sequences in concentrated solution and in the cell. Annu. Rev. Biophys. Biophys. Chem. 16, 175–204.
Figure 14–16  Regulation of the flux through multistep pathways occurs at steps that are enzyme-limited. At each of these steps (orange arrows), which are generally exergonic, the substrate is not in equilibrium with the product because the enzyme-catalyzed reaction is relatively slow. The substrate for this reaction tends to accumulate, just as river water accumulates behind a dam. In the substrate-limited reactions (blue arrows) the substrate and product are essentially at their equilibrium concentrations. At the steady state, all of the reactions in the sequence occur at the same rate, which is determined by the rate-limiting step.

Although not at equilibrium with their surroundings, adult organisms generally exist in a steady state. A constant influx of fuel and nutrients and a constant release of energy and waste products allow the organism to maintain a constant composition. When the steady state is disturbed by some change in external circumstances or fuel supply, the temporarily altered fluxes through individual metabolic pathways trigger regulatory mechanisms intrinsic to each pathway. The net effect of all of these adjustments is to return the organism to the steady state – to achieve homeostasis. Because of the central role of ATP in cellular activities, evolution has produced catabolic enzymes with regulatory properties that ensure a high steady-state concentration of ATP, “high” in this context meaning high relative to the breakdown products ADP and AMP.

The flux through a biochemical pathway depends on the activities of the enzymes that catalyze each reaction. For some of the enzymes in a pathway such as glycolysis, the reaction is essentially at equilibrium within the cell; the activity of such an enzyme is sufficiently high that the substrate is converted to product as fast as the substrate is supplied. The flux through this step is essentially substrate-limited – determined by the instantaneous concentration of the substrate.

Other cellular reactions are far from equilibrium. In the glycolytic pathway, the equilibrium constant (Keq) for the reaction catalyzed by phosphofructokinase-1 is about 250, but the mass action ratio [fructose-1,6-bisphosphate][ADP]/[fructose-6-phosphate][ATP] in a typical cell in the steady state is about 0.04. (The intracellular concentrations of some glycolytic enzymes and reactants are given in Table 14–2.)

The reaction is so far from equilibrium because the rate of conversion of fructose-6-phosphate to fructose-1,6-bisphosphate is limited by the activity of PFK-1. Increased production of fructose-6-phosphate by the preceding enzymes in the glycolytic pathway does not increase the flux through this step, but instead leads to the accumulation of the substrate, fructose-6-phosphate. Thus PFK-1 functions as a valve, regulating the flow of carbon through glycolysis; increasing the activity of this enzyme (by allosteric activation, for example) increases the overall flux through the pathway. Metabolite flux through this pathway is determined not by mass action (by substrate and product concentrations) but by how far this enzymatic valve is “opened”.

In every metabolic pathway there is at least one reaction that, in the cell, is far from equilibrium because of the relatively low activity of the enzyme that catalyzes it (Fig. 14–16). The rate of this reaction is not limited by substrate availability, but only by the activity of the enzyme. The reaction is therefore said to be enzyme-limited, and because its rate limits the rate of the whole reaction sequence, the step is called the rate-limiting step in the pathway. In general, these ratelimiting steps are very exergonic reactions and are therefore essentially irreversible under cellular conditions. Enzymes that catalyze these exergonic, rate-limiting steps are commonly the targets of metabolic regulation. In addition to very rapid allosteric enzyme regulation within individual cells, multicellular organisms use hormonal signals to coordinate the metabolic activities of different tissues and organs (Chapter 22). Hormone action alters the activities of key enzymes, often within seconds or minutes. When external circumstances change on a longer time scale, as when a human’s diet shifts from primarily fat to primarily carbohydrate, adjustments in the flux through specific pathways are brought about by changes in the number of molecules of specific regulatory enzymes. This is accomplished by changing the relative rates of synthesis and degradation of the enzymes (Chapters 26 and 27).

Many regulatory enzymes are situated at critical branch points in metabolism; their activities determine the allocation of a metabolite to each of the several pathways through which it might pass. For example, glucose-6-phosphate can be metabolized either by glycolysis or by the pentose phosphate pathway (described later in this chapter). The first enzyme unique to each of these pathways (phosphofructokinase-1 and glucose-6-phosphate dehydrogenase, respectively) catalyzes the “committed” step for its pathway. Both are regulatory enzymes, which respond to a variety of allosteric regulators that signal the need for the products of each pathway.

Cells commonly have the enzymatic capacity to carry out both the catabolism of some complex molecule into a simpler product and the anabolic conversion of that product back into the starting molecule. Glycolysis degrades glucose to pyruvate; gluconeogenesis converts pyruvate to glucose. Paired catabolic and anabolic pathways often employ many of the same enzymes – those that catalyze readily reversible reactions. Phosphoglycerate mutase, for example, acts in both glycolysis and gluconeogenesis. However, paired pathways almost invariably employ at least one reaction in the catabolic direction different from the corresponding step in the anabolic direction, and catalyzed by a different enzyme. These distinctive enzymes are the points of regulation of the two opposing pathways. The reactions catalyzed by these path-specific enzymes are generally exergonic reactions, irreversible under cellular conditions, and out of equilibrium in the steady state; they are enzyme-limited, not substrate-limited. Having separate enzymes for catabolic and anabolic pathways allows separate regulation of the flux in each direction, avoiding the wasteful “futile cycling” that would result if the breakdown and energy-consuming resynthesis of a compound were allowed to proceed simultaneously.

The regulatory enzymes that control the rate of breakdown of carbohydrates via glycolysis illustrate these general principles of metabolic regulation. Glucose catabolism is doubtless regulated in all organisms, but the regulatory mechanisms have been studied especially well in vertebrate muscle and liver.

In muscle the end served by glycolysis is ATP production, and the rate of glycolysis increases as muscle contracts more vigorously or more frequently, demanding more ATP. The liver has a different role in whole-body metabolism, and glucose metabolism in the liver is correspondingly different. The liver serves to keep a constant level of glucose in the blood, producing and exporting glucose when the tissues demand it, and importing and storing glucose when it is provided in excess in the diet.

In skeletal muscle cells (myocytes), the mobilization of stored glycogen to provide fuel for glycolysis is brought about by glycogen phosphorylase, which degrades glycogen to glucose-1-phosphate (Fig. 14–11). The case of glycogen phosphorylase is an especially instructive example of enzyme regulation. It was the first enzyme shown to be allosterically regulated and the first shown to be controlled by reversible phosphorylation. It is also one of only a few allosteric enzymes for which the detailed three-dimensional structures of the active and inactive forms are known from x-ray crystallographic studies.

In skeletal muscle, glycogen phosphorylase occurs in two forms: a catalytically active form, phosphorylase a, and a usually inactive form, phosphorylase b (Fig. 14–17); the latter predominates in resting muscle. The rate of glycogen breakdown in muscle depends in part on the ratio of phosphorylase a (active) to phosphorylase b (less active), which is adjusted by the action of hormones such as epinephrine. Phosphorylase a consists of two identical subunits (Mr 94,500), in each of

which the Ser residue at position 14 is phosphorylated. Phosphorylase b is structurally identical except that the Ser14 residues are not phosphorylated. Phosphorylase a is converted into the less active phosphorylase b by dephosphorylation, catalyzed by phosphorylase a phosphatase (Fig. 14–17). Phosphorylase b is converted back into phosphorylase a by the enzyme phosphorylase b kinase, which catalyzes phosphate transfer from ATP to Ser14.

Figure 14–17  Covalent and allosteric regulation of glycogen phosphorylase in muscle. (a) The enzyme has two identical subunits, each of which can be phosphorylated by phosphorylase b kinase at Ser14 to give phosphorylase a, a reaction promoted by Ca2+. Phosphorylase a phosphatase, also called phosphoprotein phosphatase-1, removes these phosphate groups, inactivating the enzyme. Phosphorylase b can also be activated by noncovalent binding of AMP at its allosteric sites. Conformational changes in the enzyme are indicated schematically. Liver glycogen phosphorylase undergoes similar a and b interconversions, but has different regulatory mechanisms. (b) The three-dimensional structure of the enzyme from muscle. The two subunits (gray and blue) of the glycogen phosphorylase a dimer, showing the location of the phosphates (orange) attached to the Ser14 residues (red) in each. In phosphorylase b, the amino-terminal peptide containing Ser14 is disordered. However, with the attachment of the negatively charged phosphate group at Ser14 this peptide folds toward several nearby (positively charged) Arg residues (dark blue), forcing compensatory changes in regions distant from Ser14 and activating the enzyme. AMP, the allosteric activator of phosphorylase b, binds at a site (magenta) very near Ser14. On the back side of the enzyme is a deep channel that admits the substrate glycogen to the active site, which is 3.3 nm away from the allosteric site. (c) A close-up view of the region around the phospho-Ser residue; note its proximity to the interface between dimers.

Figure 14–18  Hormonal regulation of glycogen phosphorylase in muscle and liver. A cascade of enzymatic activations leads to activation of glycogen phosphorylase by epinephrine in muscle and by glucagon in liver. When catalysts activate catalysts large amplifications of the initial signal result.
Hormones ultimately regulate the interconversion of phosphorylase a and b by regulating the activities of phosphorylase a phosphatase and phosphorylase b kinase. Epinephrine is released into the blood by the adrenal gland when an animal is suddenly confronted by a situation that requires vigorous muscular activity. Epinephrine is a signal to skeletal muscle to turn on the processes that lead to production of ATP, which will be needed for muscle contraction. Glycogen phosphorylase is activated to provide glucose-1-phosphate to be fed into the glycolytic pathway. By the cascade of events shown in Figure 14–18, the binding of epinephrine to its specific receptor in the plasma membrane of a muscle cell activates phosphorylase b kinase and inactivates phosphorylase a phosphatase, tipping the balance toward formation of the active (a) form of glycogen phosphorylase. The cascade of activations allows one molecule of hormone to cause activation of many molecules of target enzyme (glycogen phosphorylase).
When the emergency is over, release of epinephrine ceases, the phosphorylase b kinase reverts to its original, lower activity, and the ratio of phosphorylase a to phosphorylase b returns to that in resting muscle.

Superimposed on the hormonal control is faster, allosteric regulation of glycogen phosphorylase b by ATP and AMP. Phosphorylase b, the relatively inactive form, is activated by its allosteric effector AMP (Fig. 14–17), which increases in concentration in muscle during the ATP breakdown accompanying contraction. The stimulation of phosphorylase b by AMP can be prevented by high concentrations of ATP, which blocks the AMP binding site. The activity of phosphorylase b thus reflects the ratio of AMP to ATP. Phosphorylase a, which is not stimulated by AMP, is sometimes referred to as the AMP-independent form, and phosphorylase b as the AMP-dependent form.

In resting muscle nearly all the phosphorylase is in the b form, which is inactive because ATP is present at a much higher concentration than AMP. Vigorous muscular activity increases the AMP : ATP ratio, very rapidly activating (in milliseconds) phosphorylase b by allosteric means. On a longer time scale (seconds to minutes) hormone-triggered phosphorylation of phosphorylase b converts it into phosphorylase a, the activity of which is independent of the AMP : ATP ratio.

There is yet a third type of control on glycogen phosphorylase in skeletal muscle. Calcium, the intracellular signal for muscle contraction, is also an allosteric activator of phosphorylase b kinase. When a transient rise in intracellular Ca2+ triggers muscle contraction, it also accelerates conversion of phosphorylase b to the more active phosphorylase a (Fig. 14–17).
Figure 14–19  Glycogen phosphorylase as a glucose sensor. Glucose binding to an allosteric site in liver glycogen phosphorylase a induces a conformational change that exposes the phosphorylated Ser14 residues to the action of phosphorylase a phosphatase, which converts phosphorylase a to b, reducing its activity in response to high blood glucose.

The glycogen phosphorylase of liver is similar to that of muscle; it too is a dimer of identical subunits, and it undergoes phosphorylation and dephosphorylation on Ser14, interconverting the b and a forms. However, its regulatory properties are slightly different from those of the muscle enzyme, reflecting the different role of glycogen breakdown in liver. Liver glycogen serves as a reservoir that releases glucose into the blood when blood glucose levels fall below the normal level (4 to 5 mM). Glucose-1-phosphate formed by liver phosphorylase is converted (as in muscle) into glucose-6-phosphate by the action of phosphoglucomutase (p. 422). Then glucose-6-phosphatase, an enzyme present in liver but not in muscle, removes the phosphate:

Glucose-6-phosphate + H2O   →   glucose + Pi

When the blood glucose level is low, the free glucose produced from glycogen in the liver by these reactions is released into the bloodstream and carried to tissues (such as the brain) that require it as a fuel. (For most tissues, glucose is only one of several equally useful fuels.)

Glycogen phosphorylase of liver, like that of muscle, is under hormonal control. Glucagon is a hormone released by the pancreas when blood glucose levels fall below normal. When glucagon binds to its receptor in the plasma membrane of a hepatocyte, a cascade of events essentially similar to that in muscle (Fig. 14–18) results in the conversion of phosphorylase b to phosphorylase a, increasing the rate of glycogen breakdown and thereby increasing the rate of glucose release into the blood.

Liver glycogen phosphorylase, like that of muscle, is subject to allosteric regulation, but in this case the allosteric regulator is glucose, not AMP. When the concentration of glucose in the blood rises, glucose enters hepatocytes and binds to the regulatory site of glycogen phosphorylase a, causing a conformational change that exposes the phosphorylated Ser14 residues to dephosphorylation by phosphorylase a phosphatase (Fig. 14–19). In this way, glycogen phosphorylase a acts as the glucose sensor of liver, slowing the breakdown of glycogen whenever the level of blood glucose is high.

Hexokinase, which catalyzes the entry of free glucose into the glycolytic pathway, is another regulatory enzyme. The hexokinase of myocytes has a high affinity for glucose (it is half saturated at about 0.1 mM). Glucose entering myocytes from the blood (in which the glucose concentration is 4 to 5 mM) produces an intracellular glucose concentration high enough to saturate hexokinase, so that it normally acts at its maximal rate. Muscle hexokinase is allosterically inhibited by its product, glucose-6-phosphate. Whenever the concentration of glucose6-phosphate in the cell rises above its normal level, hexokinase is temporarily and reversibly inhibited, bringing the rate of glucose-6-phosphate formation into balance with the rate of its utilization and reestablishing the steady state.

Mammals have several forms of hexokinase, all of which catalyze the conversion of glucose into glucose-6-phosphate. Different proteins that catalyze the same reaction are called isozymes (Box 14–3). The predominant hexokinase isozyme in liver is hexokinase D, also called glucokinase, which differs in two important respects from the hexokinase isozymes in muscle.

First, the glucose concentration at which glucokinase is half-saturated (about 10 mM) is higher than the usual concentration of glucose in the blood. Because the concentration of glucose in liver is maintained at a level close to that in the blood by an efficient glucose transporter, this property of glucokinase allows its direct regulation by the level of blood glucose. When the glucose concentration in the blood is high, as it is after a meal rich in carbohydrates, excess blood glucose is transported into hepatocytes, where glucokinase converts it into glucose-6-phosphate.

Second, glucokinase is inhibited not by its reaction product glucose-6-phosphate but by its isomer, fructose-6-phosphate, which is always in equilibrium with glucose-6-phosphate because of the action of phosphoglucose isomerase. The partial inhibition of glucokinase by fructose-6-phosphate is mediated by an additional protein, the regulator protein. This regulator protein also has affinity for fructose-1-phosphate, which competes with fructose-6-phosphate and cancels its inhibitory effect on glucokinase. Because fructose-1-phosphate is present

in liver only when there is fructose in the blood, this property of the regulator protein explains the observation that ingested fructose stimulates the phosphorylation of glucose in the liver.

The pancreatic β cells, which are responsible for the release of insulin when blood glucose levels rise above normal, also contain glucokinase and the inhibitory regulator protein.

B O X  14–3
Isozymes: Different Proteins That Catalyze the Same Reaction

The several forms of hexokinase found in mammalian tissues are but one example of a common situation in which the same reaction is catalyzed by two or more different molecular forms of an enzyme. These multiple forms, called isozymes or isoenzymes, may occur in the same species, in the same tissue, or even in the same cell. The different forms of the enzyme generally differ in kinetic or regulatory properties, in the form of cofactor they use (NADH or NADPH for dehydrogenase isozymes, for example) or in their subcellular distribution (soluble or membrane-bound). Isozymes commonly have similar, but not identical, amino acid sequences, and in many cases they clearly share an evolutionary origin.

One of the first enzymes found to have isozymes was lactate dehydrogenase (LDH) (p. 416). LDH occurs in vertebrate tissues as at least five different isozymes separable by electrophoresis. All LDH isozymes contain four polypeptide chains (each of Mr 33,500), but the five isozymes contain different ratios of two kinds of polypeptides that differ in composition and sequence. The A chains (also designated M for muscle) and the B chains (also designated H for heart) are encoded by two different genes. In skeletal muscle the predominant isozyme contains four A chains, and in heart the predominant isozyme contains four B chains. LDH isozymes in other tissues are a mixture of the five possible forms, which may be designated A4, A3B, A2B2, AB3, and B4. The different LDH isozymes have significantly different values of Vmax and Km, particularly for pyruvate. The properties of LDH isozyme A4 favor rapid reduction of very low concentrations of pyruvate to lactate in skeletal muscle, whereas those of isozyme B4 tend to favor rapid oxidation of lactate to pyruvate in the heart.

The distribution of different isozyme forms of a given enzyme reflects at least four factors:

1. The differing metabolic patterns in different organs. The two forms of glycogen phosphorylase found in skeletal muscle and in liver differ in their regulatory properties, reflecting the different roles of glycogen breakdown in these two tissues, as described in the text.

2. The different locations and metabolic roles of a given enzyme within one type of cell. The isocitrate dehydrogenase isozymes of the cytosol and the mitochondrion are an example (Chapter 15).

3. The differentiation and development of adult tissues from their embryonic or fetal forms. For example, the fetal liver has a characteristic isozyme distribution of LDH, which changes as the organ undergoes differentiation to its adult form. An interesting discovery is that some of the enzymes of glucose catabolism in malignant (cancer) cells occur as their fetal, not adult, isozymes.

4. The fine-tuning of metabolic rates through the different responses of isozyme forms to allosteric modulators. Hexokinase D (glucokinase) of liver and the hexokinase isozymes found in other tissues differ in their sensitivity to inhibition by their product, glucose-6-phosphate (p. 432).

In vertebrates there are at least three isozymes of pyruvate kinase, differing somewhat in their tissue distribution and in their response to modulators. High concentrations of ATP inhibit pyruvate kinase allosterically, by decreasing the affinity of the enzyme for its substrate phosphoenolpyruvate (PEP). The level of PEP normally found in cells is not high enough to saturate the enzyme, and the reaction rate will accordingly be low at normal PEP concentrations.

Pyruvate kinase is also inhibited by acetyl-CoA and by long-chain fatty acids, both important fuels for the citric acid cycle. (Recall that acetyl-CoA (acetate) is produced by the catabolism of fats and amino acids, as well as by glucose catabolism; see Fig. 4a, p. 362.) Because the citric acid cycle is a major source of energy for ATP production, the availability of these other fuels reduces the dependence on glycolysis for ATP.

Thus, whenever the cell has a high concentration of ATP, or whenever ample fuels are already available for energy-yielding respiration, glycolysis is inhibited by the slowed action of pyruvate kinase. When the ATP concentration falls, the affinity of pyruvate kinase for PEP increases, enabling the enzyme to catalyze ATP synthesis even though the concentration of PEP is relatively low. The result is a high steady-state concentration of ATP.

Figure 14–20  (a) A ribbon diagram of E. coli phosphofructokinase-1, showing two of its four identical subunits. Each subunit has its own catalytic site and its own binding sites for the allosteric activators. (b) Allosteric regulation of muscle PFK-1 by ATP, shown by a substrate-activity curve. At low concentrations of ATP the K0.5 (p. 231) for fructose 6-phosphate is relatively low, enabling the enzyme to function at a high rate at relatively low concentrations of fructose-6-phosphate. At high ATP, K0.5 for fructose-6-phosphate is greatly increased, as indicated by the sigmoid relationship between substrate concentration and enzyme activity. (c) A summary of the regulators affecting PFK-1 activity.

Glucose-6-phosphate can flow either into glycolysis or through one of the secondary oxidative pathways described later in this chapter. The irreversible reaction catalyzed by PFK-1 is the step that commits a cell to the passage of glucose through glycolysis. In addition to the binding sites for its substrates, fructose-6-phosphate and ATP, this complex enzyme has several regulatory sites where allosteric activators or inhibitors bind.

ATP is not only the substrate for PFK-1, but also the end product of the glycolytic pathway. When high ATP levels signal that the cell is producing ATP faster than it is consuming it, ATP inhibits PFK-1 by binding to an allosteric site and lowering the affinity of the enzyme for its substrate fructose-6-phosphate (Fig. 14–20). ADP and AMP, which rise in concentration when the consumption of ATP outpaces its production, act allosterically to relieve this inhibition by ATP. These effects combine to produce higher enzyme activity when fructose-6-phosphate, ADP, or AMP builds up, and lower activity when ATP accumulates.

Citrate (the ionized form of citric acid), a key intermediate in the aerobic oxidation of pyruvate (Chapter 15), also serves as an allosteric regulator of PFK-1; high citrate concentration increases the inhibitory effect of ATP, further reducing the flow of glucose through glycolysis. In this case, as in several others to be encountered later, citrate serves as an intracellular signal that the cell's needs for energy-yielding metabolism and for biosynthetic intermediates are being met.

The most significant allosteric regulator of PFK-1 is fructose-2,6-bisphosphate, which, as noted earlier, strongly activates the enzyme. The concentration of fructose-2,6-bisphosphate in liver decreases in response to the hormone glucagon, slowing glycolysis and stimulating glucose synthesis in liver.
Figure 14–21  The reaction in gluconeogenesis that bypasses the irreversible phosphofructokinase-1 reaction in glycolysis. The conversion of fructose-1,6-bisphosphate to fructose-6-phosphate is catalyzed by fructose-1,6-bisphosphatase (called FBPase-1 to distinguish it from a similar enzyme described in Chapter 19).

Most organisms can synthesize glucose from simpler precursors such as pyruvate or lactate. In mammals this process, called gluconeogenesis, occurs primarily in the liver, and its role is to provide glucose for export to other tissues when other sources of glucose are exhausted. Gluconeogenesis employs most of the same enzymes that act in glycolysis, but it is not simply the reversal of glycolysis. Seven of the glycolytic reactions are freely reversible, and the enzymes that catalyze these reactions also function in gluconeogenesis. Three reactions of glycolysis are so exergonic as to be essentially irreversible: those catalyzed by hexokinase, phosphofructokinase-1, and pyruvate kinase. Detours around each of these irreversible steps are employed in gluconeogenesis. For example, in gluconeogenesis the conversion of fructose-1,6-bisphosphate to fructose-6-phosphate is catalyzed by fructose-1,6-bisphosphatase (FBPase-1) (Fig. 14–21).

To prevent futile cycling in which glucose is simultaneously degraded by glycolysis and resynthesized by gluconeogenesis (Chapter 19), the enzymes unique to each pathway are reciprocally regulated by common allosteric effectors. Fructose-2,6-bisphosphate, a potent activator of liver PFK-1 and therefore of glycolysis, also inhibits FBPase-1, thereby slowing gluconeogenesis.

Glucagon, the hormone that signals low blood sugar, lowers the level of fructose-2,6-bisphosphate in liver, slowing the consumption of glucose by glycolysis and stimulating the production of glucose for export by gluconeogenesis. We will return to a more complete discussion of this coordinate regulation in Chapter 19, when we have discussed gluconeogenesis in more detail.

Fructose-2,6-bisphosphate is found in all animals, in fungi, and in some plants, but not in bacteria. It stimulates all known animal PFK-1 activities as well as PFK-1 from yeast. In plants, fructose-2,6-bisphosphate also regulates carbohydrate metabolism, but by mechanisms not identical with those in liver; plants do not, of course, have glucagon. Fructose-2,6-bisphosphate activates the PPi-dependent phosphofructokinase of plants that is responsible for fructose-1,6-bisphosphate formation in glycolysis (p. 407), but does not activate the ATP-dependent PFK-1 of plants. Plant PFK-1 is, however, strongly inhibited by phosphoenolpyruvate, a glycolytic intermediate downstream from fructose-1,6-bisphosphate.

In animal tissues, most of the glucose consumed is catabolized via glycolysis to pyruvate. Most of the pyruvate in turn is oxidized via the citric acid cycle. The main function of glucose catabolism by this route is to generate ATP. There are, however, other catabolic pathways taken by glucose that lead to specialized products needed by the cell, and these pathways constitute part of the secondary metabolism of glucose. Two such pathways produce pentose phosphates and uronic and ascorbic acids.

Figure 14–22  The oxidative reactions of the pentose phosphate pathway, leading to D-ribose-5-phosphate and producing NADPH.

The pentose phosphate pathway, also called the phosphogluconate pathway (Fig. 14–22), produces NADPH and ribose-5-phosphate. Recall that NADPH is a carrier of chemical energy in the form of reducing power (Chapter 13). In mammals this function is especially prominent in tissues actively carrying out the biosynthesis of fatty acids and steroids from small precursors, particularly the mammary gland, adipose tissue, the adrenal cortex, and the liver. The biosynthesis of fatty acids requires reducing power in the form of NADPH to reduce the double bonds and carbonyl groups of intermediates in this process (Chapter 20). Other tissues less active in synthesizing fatty acids, such as skeletal muscle, are virtually lacking in the pentose phosphate pathway. A second function of the pentose phosphate pathway is to generate essential pentoses, particularly D-ribose, used in the biosynthesis of nucleic acids (Chapter 21).

The first reaction of the pentose phosphate pathway is the enzymatic dehydrogenation of glucose-6-phosphate by glucose-6-phosphate dehydrogenase to form 6-phosphoglucono-δ-lactone, an intramolecular ester, which is hydrolyzed to the free acid 6-phosphogluconate by a specific lactonase (Fig. 14–22). NADP+ is the electron acceptor, and the overall equilibrium lies far in the direction of formation of NADPH. In the next step 6-phosphogluconate undergoes dehydrogenation and decarboxylation by 6-phosphogluconate dehydrogenase to form the ketopentose D-ribulose-5-phosphate, a reaction that generates a second molecule of NADPH. Phosphopentose isomerase then converts D-ribulose-5-phosphate into its aldose isomer D-ribose-5-phosphate. In some tissues, the pentose phosphate pathway ends at this point, and its overall equation is then written

Glucose-6-phosphate + 2NADP+ + H2O   →   ribose-5-phosphate + CO2 + 2NADPH + 2H+

The net result is the production of NADPH for reductive biosynthetic reactions and the production of ribose-5-phosphate as a precursor for nucleotide synthesis.

Figure 14–23  (a) The nonoxidative reactions of the pentose phosphate pathway convert pentose phosphates back into hexose phosphates, allowing the oxidative reactions (see Fig. 14–22) to continue. The enzymes transaldolase and transketolase (discussed in more detail in Chapter 19) are specific to this pathway; the other enzymes also serve in the glycolytic or gluconeogenic pathways. (b) A simplified schematic diagram showing the pathway leading from six pentoses (5C) to five hexoses (6C). Note that this involves two sets of the interconversions shown in (a).
In tissues that require primarily NADPH rather than ribose-5phosphate, pentose phosphates are recycled into glucose-6-phosphate in a series of reactions (Fig. 14–23) that will be examined in more detail in Chapter 19. First, ribulose-5-phosphate is epimerized to xylulose-5-phosphate. Then, in a series of rearrangements of the carbon skeletons of sugar phosphate intermediates, six five-carbon sugar phosphates are converted into five six-carbon sugar phosphates (Fig. 14–23b), completing the cycle and allowing continued oxidation of glucose-6-phosphate with the production of NADPH.
In the nonoxidative part of the pentose phosphate pathway (Fig. 14–23a), transketolase, a thiamine pyrophosphate-dependent enzyme, catalyzes the transfer of a two-carbon fragment (C-1 and C-2) of xylulose-5-phosphate to ribose-5-phosphate, forming the seven-carbon product sedoheptulose-7-phosphate; the remaining three-carbon fragment of xylulose is glyceraldehyde-3-phosphate. (The detailed mechanism for transketolase is shown in Fig. 19–25.) Transaldolase then catalyzes a reaction similar to the aldolase reaction in glycolysis: a three-carbon fragment is removed from sedoheptulose-7-phosphate and condensed with glyceraldehyde-3-phosphate, forming fructose-6-phosphate; the remaining four-carbon fragment of sedoheptulose is erythrose-4-phosphate. Now transketolase acts again, forming fructose-6-phosphate and glyceraldehyde-3-phosphate from erythrose-4–phosphate and xylulose-5-phosphate. Two molecules of glyceraldehyde-3-phosphate formed by two iterations of these reactions can be converted into fructose-1,6-bisphosphate (Fig. 14–23b). The cycle is then complete: six pentose phosphates have been converted back into five hexose phosphates.

All of the reactions of the nonoxidative part of the pentose phosphate pathway are readily reversible, and thus also provide a means of converting hexose phosphates into pentose phosphates. As we shall see in Chapter 19, this is essential in the fixation of CO2 by photosynthetic plants.

Another secondary pathway for glucose leads to two specialized products: D-glucuronate, important in the detoxification and excretion of foreign organic compounds, and L-ascorbic acid or vitamin C. Although the amount of glucose diverted into this secondary pathway is very small compared with the large amounts of glucose proceeding through glycolysis and the citric acid cycle, the products are vital to the organism.

In this pathway (Fig. 14–24) glucose-1-phosphate is first converted into UDP-glucose by reaction with UTP. The glucose portion of UDP-glucose is then dehydrogenated to yield UDP-glucuronate, another example (see also Fig. 14–13) of the use of UDP derivatives as intermediates in the enzymatic transformations of sugars.

UDP-glucuronate is the glucuronosyl donor used by a family of detoxifying enzymes that act on a variety of relatively nonpolar drugs, environmental toxins, and carcinogens. The conjugation of these compounds with glucuronate (glucuronidation) converts them into much more polar derivatives that are more easily cleared from the blood by the kidneys and excreted in the urine. For example, the sedative drug phenobarbital, the anti-AIDS drug AZT, and the hydroxylated form of the carcinogen benzo[a]pyrene (3-hydroxybenzo[a]pyrene) all undergo glucuronidation catalyzed by UDP-glucuronosyl transferases in the human liver (Fig. 14–25). Chronic exposure to the drug or toxin induces increased synthesis of the enzyme specific for that compound, increasing tolerance for the drug or resistance to the toxin. UDP-glucuronate is also the precursor of the glucuronate residues of such acidic polysaccharides as hyaluronate and chondroitin sulfate (see Fig. 11–20).
Figure 14–25  Detoxification of 3-hydroxybenzo[a]pyrene, a toxic component of tobacco smoke. Glucuronidation by transfer of glucuronate from UDP-glucuronate converts the nonpolar toxin to a polar compound more easily removed by the kidneys.
D-Glucuronate is an intermediate in the conversion of D-glucose into L-ascorbic acid (Fig. 14–24). It is reduced by NADPH to the sixcarbon sugar acid L-gulonate, which is converted into its lactone. L-Gulonolactone then undergoes dehydrogenation by the flavoprotein gulonolactone oxidase to yield L-ascorbic acid. Some animal species, including humans, guinea pigs, monkeys, some birds, and some fish lack the enzyme gulonolactone oxidase and are unable to synthesize ascorbic acid; they require it ready-made in the diet (as vitamin C).

Humans who do not obtain enough vitamin C in the diet develop the serious disease scurvy, in which the synthesis of connective tissue containing collagen is defective. The symptoms of scurvy include swollen and bleeding gums with loosened teeth, stiffness and soreness of joints, bleeding under the skin, and slow wound healing. For centuries the disease was very common among sailors on long sea voyages, during which no fresh fruit was available, and in 1753 the Scottish naval surgeon James Lind showed that scurvy was prevented and cured by ingestion of citrus juice. In 1932 the antiscurvy vitamin C was isolated from lemon juice and named ascorbic acid (from the Latin scorbutus, meaning “scurvy”).
Figure 15–1  Catabolism of proteins, fats, and carbohydrates occurs in the three stages of cellular respiration. Stage 1: Oxidation of fatty acids, glucose, and some amino acids yields acetyl-CoA. Stage 2: Oxidation of acetyl groups via the citric acid cycle includes four steps in which electrons are abstracted. Stage 3: Electrons carried by NADH and FADH2 are funneled into a chain of mitochondrial (or plasma membrane-bound, in bacteria) electron carriers – the respiratory chain – ultimately reducing O2 to H2O. This electron flow drives the synthesis of ATP, in the process of oxidative phosphorylation.
Chapter 15
The Citric Acid Cycle

We saw in Chapter 14 how cells obtain energy (ATP) by fermentation, breaking down glucose in the absence of oxygen. However, most eukaryotic cells and many bacteria normally are aerobic and oxidize their organic fuels completely to CO2 and H2O. Under these conditions the pyruvate formed in the glycolytic breakdown of glucose is not reduced to lactate, ethanol, or some other fermentation product, as occurs under anaerobic conditions, but instead is oxidized to CO2 and H2O in the aerobic phase of catabolism, respiration. In the broader physiological or macroscopic sense, respiration refers to a multicellular organism’s uptake of O2 from its environment and release of CO2, but biochemists and cell biologists use the term in a microscopic sense to refer to the molecular processes involved in O2 consumption and CO2 formation by cells. This latter process may be more precisely termed cellular respiration.

Cellular respiration occurs in three major stages (Fig. 15–1). In the first stage, organic fuel molecules – glucose, fatty acids, and some amino acids – are oxidized to yield two-carbon fragments in the form of the acetyl group of acetyl-coenzyme A (acetyl-CoA). In the second stage, these acetyl groups are fed into the citric acid cycle, which enzymatically oxidizes them to CO2. The energy released by oxidation is conserved in the reduced electron carriers NADH and FADH2. In the third stage of respiration, these reduced cofactors are themselves oxidized, giving up protons (H+) and electrons. The electrons are transferred along a chain of electron-carrying molecules, known as the respiratory chain, to O2, which they reduce to form H2O. During this process of electron transfer much energy is released and conserved in the form of ATP, in the process called oxidative phosphorylation. Respiration is more complex than glycolysis and is believed to have evolved much later, after the appearance of cyanobacteria, which added oxygen – the electron acceptor of respiration – to the earth’s atmosphere.

We will discuss the third stage of cellular respiration (electron transfer and oxidative phosphorylation) in Chapter 18. In this chapter we examine the complete oxidation of pyruvate and the citric acid cycle, also called the tricarboxylic acid cycle or the Krebs cycle (after its discoverer). We consider first the cycle reactions and the enzymes that catalyze them. Because intermediates of the citric acid cycle are often used as biosynthetic precursors, some means of replenishing these intermediates is essential to the continued operation of the cycle; we discuss several such replenishing reactions. The mechanisms

that regulate the flux of material through the citric acid cycle are then considered. Finally, we describe a metabolic sequence, the glyoxylate pathway, that employs some of the same enzymes and reactions that occur in the citric acid cycle to enable the net synthesis of glucose from stored triacylglycerols.

In aerobic organisms, glucose and other sugars, fatty acids, and most of the amino acids are ultimately oxidized to CO2 and H2O via the citric acid cycle. Before they can enter the cycle, the carbon skeletons of sugars and fatty acids must be degraded to the acetyl group of acetyl-CoA, the form in which the citric acid cycle accepts most of its fuel input. Many amino acid carbons also enter the cycle this way, although several amino acids are degraded to other intermediates of the cycle. In Chapters 16 and 17, respectively, we shall see how fatty acids and amino acids enter the citric acid cycle. Here we consider how pyruvate, derived from glucose by glycolysis, is oxidized to yield acetyl-CoA and CO2 by a structured cluster of three enzymes, the pyruvate dehydrogenase complex, located in the mitochondria of eukaryotic cells and in the cytosol of prokaryotes.

A careful examination of this enzyme complex can be rewarding in several respects. The pyruvate dehydrogenase complex is a classic example, very well studied, of a multienzyme complex in which a series of chemical intermediates remain bound to the surface of the enzyme molecules as the substrate is transformed into the imal product. Five cofactors participate in this key reaction mechanism, all of which are coenzymes derived from vitamins. The regulation of this enzyme complex also illustrates how a combination of covalent modification and allosteric regulation results in precisely regulated flux through a metabolic step. Finally, the pyruvate dehydrogenase complex is the prototype for two other important enzyme complexes that we will encounter: α-ketoglutarate dehydrogenase (of the citric acid cycle) and the branched-chain α-ketoacid dehydrogenase involved in the oxidative degradation of several amino acids (Chapter 17). The remarkable similarity in the protein structure, cofactor requirements, and reaction mechanisms of these three complexes doubtless reflects a common evolutionary origin.

Figure 15–2  The overall reaction catalyzed by the pyruvate dehydrogenase complex. The five coenzymes involved in this reaction, and the three enzymes that make up the enzyme complex, are discussed in the text.

The overall reaction catalyzed by the pyruvate dehydrogenase complex is oxidative decarboxylation, an irreversible oxidation process in which the carboxyl group is removed from pyruvate as a molecule of CO2 and the two remaining carbons become the acetyl group of acetyl-CoA (Fig. 15–2). The NADH formed in this reaction gives up a hydride ion with its two electrons ( ⁚ H) to the respiratory chain (Fig. 15–1), which carries the electrons to oxygen or, in anaerobic microorganisms, to an alternative electron acceptor such as sulfur. This electron transfer to oxygen ultimately generates three molecules of ATP per pair of electrons.

The irreversibility of the pyruvate dehydrogenase reaction has been proved by isotopic labeling experiments: radioactively labeled CO2 cannot be reattached to acetyl-CoA to yield pyruvate labeled in the carboxyl group.

Figure 15–3  The structure of coenzyme A. A hydroxyl group of pantothenic acid is joined to a modified ADP moiety by a phosphate ester bond, and its carboxyl group is attached to β-mercaptoethylamine in amide linkage. The hydroxyl group at the 3′ position of the ADP moiety has a phosphate group not present in ADP itself. The –SH group of the mercaptoethylamine moiety forms a thioester with acetate in acetyl-CoA (lower left). Coenzyme A is abbreviated as CoA, and acetyl-coenzyme A as acetyl-CoA; the reactive –SH group is generally shown only in chemical structures.
Figure 15–4  Lipoic acid in amide linkage with the side chain of a Lys residue is the prosthetic group of dihydrolipoyl transacetylase (E2). The lipoyl group occurs in oxidized (disulfide) or reduced (dithiol) form and can act as a carrier of both hydrogen and an acetyl (or other acyl) group.

The combined dehydrogenation and decarboxylation of pyruvate to acetyl-CoA (Fig. 15–2) involves the sequential action of three different enzymes, as well as five different coenzymes or prosthetic groups – thiamine pyrophosphate (TPP), flavin adenine dinucleotide (FAD), coenzyme A (CoA), nicotinamide adenine dinucleotide (NAD), and lipoate. Four different vitamins required in human nutrition are vital components of this system: thiamin (in TPP), riboflavin (in FAD), niacin (in NAD), and pantothenate (in coenzyme A).

We have already described the roles of FAD and NAD as electron carriers (Chapter 13), and we have encountered TPP as the coenzyme of the enzyme that catalyzes decarboxylation of pyruvate to acetaldehyde in alcohol fermentation (see Fig. 14–9). Pantothenate, present in all living organisms, is an essential component of coenzyme A (Fig. 15–3).

Coenzyme A has a reactive thiol (–SH) group that is critical to its role as an acyl carrier in a number of metabolic reactions; acyl groups become covalently linked to this thiol group, forming thioesters. Because of their relatively high free energy of hydrolysis (see Figs. 13–6, 13–7), thioesters have a high acyl group transfer potential, donating their acyl groups to a variety of acceptor molecules. The acyl group attached to coenzyme A may thus be thought of as activated for group transfer.

The fifth cofactor for the pyruvate dehydrogenase reaction, lipoate (Fig. 15–4), has two thiol groups, both essential to its role as cofactor. In the reduced form of lipoate both sulfur atoms are present as –SH groups, but oxidation produces a disulfide (–S–S–) bond, similar to that between two Cys residues in a protein. Because of this capacity to undergo oxidation–reduction reactions, lipoate can serve both as an electron carrier and as an acyl carrier; both functions are important in the action of the pyruvate dehydrogenase complex.

Figure 15–5  (a) Electron micrograph of the pyruvate dehydrogenase complex isolated from E. coli, showing its subunit structure. (b) Interpretive model of the organization of the mammalian pyruvate dehydrogenase complex. The 24 E2 subunits are depicted as having an inner catalytic domain (green) with an attached flexible lipoyllysyl domain (red) and an E1/E3 binding domain (blue) joined by linker segments (gray). The complex also contains 24 dimeric E1 components (orange) and 6 dimeric E3 components (yellow). Note that the mammalian complex is larger and has more subunits than the E. coli complex.

The pyruvate dehydrogenase complex consists of multiple copies of each of the three enzymes pyruvate dehydrogenase (E1), dihydrolipoyl transacetylase (E2), and dihydrolipoyl dehydrogenase (E3) (Table 15–1). The number of copies of each subunit, and therefore the size of the complex, varies from one organism to another. The pyruvate dehydrogenase complex isolated from E. coli (Mr > 4.5 × 106) is about 45 nm in diameter, slightly larger than a ribosome, and can be visualized with the electron microscope (Fig. 15–5a). The “core” of the cluster, to which the other enzymes are attached, is dihydrolipoyl transacetylase (E2). In the complex from E. coli, 24 copies of this polypeptide chain, each containing three molecules of covalently bound lipoate, constitute this core. In the complex from mammals, there are 60 copies of E2 and six of a related protein, protein X, which also contains covalently bound lipoate. The attachment of lipoate to the ends of

Lys side chains in E2 produces lipoyllysyl groups (Fig. 15–4) – long, flexible arms that can carry acetyl groups from one active site to another in the pyruvate dehydrogenase complex. Bound to the core of E2 molecules are 12 copies of pyruvate dehydrogenase (E1), each composed of two identical subunits, and six copies of dihydrolipoyl dehydrogenase (E3), each also composed of two identical subunits. Pyruvate dehydrogenase (E1) contains bound TPP, and dihydrolipoyl dehydrogenase (E3) contains bound FAD. Two regulatory proteins that are also part of the pyruvate dehydrogenase complex (a protein kinase and a phosphoprotein phosphatase) will be discussed below.

Table 15–1  Subunit composition of the E. coli pyruvate
dehydrogenase complex
 Enzyme                                Coenzyme(s)      Molecular
weight of     
Number of
per complex 
       dehydrogenase (E1)
       transacetylase (E2)   
       dehydrogenase (E3)
Lipoate, CoA   

Source: Modified from Eley, M.H., Namihira, G., Hamilton, L.. Munk, P., & Reed, L.J. (1972)
α-Ketoacid dehydrogenases. XVIII: subunit composition of the E. coli pyruvate dehydrogenase complex.
Arch. Biochem. Biophys. 152, 655–669.
Figure 15–6  Steps in the oxidative decarboxylation of pyruvate to acetyl-CoA by the pyruvate dehydrogenase complex. The fate of pyruvate is traced in pink. In step pyruvate reacts with the bound thiamine pyrophosphate (TPP) of pyruvate dehydrogenase (E1), undergoing decarboxylation to form the hydroxyethyl derivative. Pyruvate dehydrogenase also carries out step , the transfer of two electrons and the acetyl group from TPP to the oxidized form of the lipoyllysyl group of the core enzyme, dihydrolipoyl transacetylase (E2), to form the acetyl thioester of the reduced lipoyl group. Step is a transesterification in which the –SH group of CoA replaces the –SH group of E2 to yield acetyl-CoA and the fully reduced (dithiol) form of the lipoyl group. In step dihydrolipoyl dehydrogenase (E3) promotes transfer of two hydrogen atoms from the reduced lipoyl groups of E2 to the FAD prosthetic group of E3, restoring the oxidized form of the lipoyllysyl group of E2 (shaded yellow). In step the reduced FADH2 group on E3 transfers a hydride ion to NAD+, forming NADH. The enzyme complex is now ready for another catalytic cycle. The structures of TPP and its hydroxyethyl derivative are shown in Fig. 14–9. E1, pyruvate dehydrogenase; E2, dihydrolipoyl transacetylase; E3, dihydrolipoyl dehydrogenase.

Figure 15–6 shows schematically how the pyruvate dehydrogenase complex carries out the five consecutive reactions in the decarboxylation and dehydrogenation of pyruvate. Step is essentially identical to the reaction catalyzed by pyruvate decarboxylase (see Fig. 14–9c); C-1 of pyruvate is released as CO2, and C-2, which in pyruvate has the oxidation state of an aldehyde, is attached to TPP as a hydroxyethyl group. In step this group is oxidized to a carboxylic acid (acetate).

The two electrons removed in the oxidation reaction reduce the –S–S– of a lipoyl group on E2 to two thiol (–SH) groups. The acetate produced in this oxidation–reduction reaction is first esterified to one of the lipoyl –SH groups, then transesterified to CoA to form acetyl-CoA (step ). Thus the energy of oxidation drives the formation of a high-energy thioester of acetate. The remainder of the reactions catalyzed by the pyruvate dehydrogenase complex (steps and ) are electron transfers necessary to regenerate the disulfide form of the lipoyl group of E2 to prepare the enzyme complex for another round of oxidation. The electrons removed from the hydroxyethyl group derived from pyruvate eventually appear in NADH after passing through FAD.

Central to the process are the swinging lipoyllysyl arms of E2, which pass the two electrons and the acetyl group derived from pyruvate from E1 to E3. All these enzymes and coenzymes are clustered, allowing the intermediates to react quickly without ever diffusing away from the surface of the enzyme complex. The five-reaction sequence shown in Figure 15–6 is another example of substrate channeling, as described in Chapter 14 (see Fig. 14–8).

As one might predict, mutations in the genes for pyruvate dehydrogenase subunits, or thiamin deficiency in the diet, have severe consequences. Thiamin-deficient animals are unable to oxidize pyruvate normally, which is of particular importance in the brain; this organ usually obtains all its energy from the aerobic oxidation of glucose, and pyruvate oxidation is therefore vital. Beriberi, a disease that results from dietary deficiency of thiamin, is characterized by loss of neural function. This disease occurs primarily in populations that depend on white (polished) rice for most of their food; white rice lacks the hulls in which most of the thiamin of rice is found. People who habitually consume large amounts of alcohol can also develop thiamin deficiency; much of their dietary intake is in the form of the “empty (vitamin-free) calories” of distilled spirits. An elevated level of pyruvate in the blood is often an indicator of defects in pyruvate oxidation due to one of these causes.
Figure 15–7  The reactions of the citric acid cycle. The carbon atoms shaded in red are those originally derived from the acetate of acetyl-CoA in the first turn of the cycle. These carbons are not the ones released as CO2 in the first turn. Note that in fumarate, the two-carbon group derived from acetate can no longer be specifically denoted; because succinate and fumarate are symmetric molecules, C-1 and C-2 are indistinguishable from C-4 and C-3. The number beside each reaction step corresponds to a numbered heading in the text. Steps , , and are essentially irreversible in the cell; all of the others are reversible.

Having seen how acetyl-CoA is formed from pyruvate, we are ready to examine the citric acid cycle. An overview of how the cycle functions will be helpful. First we note a fundamental difference between glycolysis and the citric acid cycle. Glycolysis takes place by a linear sequence of enzyme-catalyzed steps, whereas the sequence of reactions in the citric acid cycle is cyclical. To begin a turn of the cycle (Fig. 15–7), acetyl-CoA donates its acetyl group to the four-carbon compound oxaloacetate to form the six-carbon citrate. Citrate is then transformed into isocitrate, also a six-carbon molecule, which is dehydrogenated with loss of CO2 to yield the five-carbon compound aketoglutarate. The latter undergoes loss of CO2 and ultimately yields the four-carbon compound succinate and a second molecule of CO2. Succinate is then enzymatically converted in three steps into the four-carbon oxaloacetate, with which the cycle began; thus, oxaloacetate is ready to react with another molecule of acetyl-CoA to start a second turn. In each turn of the cycle, one acetyl group (two carbons) enters as acetyl-CoA and two molecules of CO2 leave. In each turn, one molecule of oxaloacetate is used to form citrate but – after a series of reactions – the oxaloacetate is regenerated. Therefore no net removal of oxaloacetate occurs; one molecule of oxaloacetate can theoretically suffice to bring about oxidation of an infinite number of acetyl groups. Four of the eight steps in this process are oxidations, in which the energy of oxidation is conserved, with high efficiency, in the formation of reduced cofactors (NADH and FADH2).

After discussing the individual reactions of the citric acid cycle in detail, we will briefly consider the experiments that led to its discovery, and the evolutionary origins of the cycle.

Although the citric acid cycle is central to energy-yielding metabolism, its role is not limited to energy conservation. Four- and five-carbon intermediates of the cycle serve as biosynthetic precursors for a wide variety of products. To replace intermediates removed for this purpose, cells employ anaplerotic (replenishing) reactions, which are briefly described.

Eugene Kennedy and Albert Lehninger showed in 1948 that in eukaryotes the entire set of reactions of the citric acid cycle takes place in the mitochondria. Isolated mitochondria were found to contain not only all the enzymes and coenzymes required for the citric acid cycle but also all the enzymes and proteins necessary for the last stage of respiration, namely, electron transfer and ATP synthesis by oxidative phosphorylation. As we shall see in later chapters, mitochondria also contain the enzymes that catalyze the oxidation of fatty acids to acetyl-CoA and the oxidative degradation of amino acids to acetyl-CoA or, for some amino acids, α-ketoglutarate, succinyl-CoA, or oxaloacetate. Thus in nonphotosynthetic eukaryotes the mitochondrion is the site of most energy-yielding oxidative reactions and of the synthesis of ATP coupled to those reactions. In photosynthetic eukaryotes, mitochondria are the major site of ATP production in the dark, but in daylight chloroplasts produce most of the ATP. Most prokaryotes contain the enzymes of the citric acid cycle in their cytosol, and their plasma membrane plays a role analogous to that of the inner mitochondrial membrane in ATP synthesis (Chapter 18).

In examining the eight successive reaction steps of the citric acid cycle, we will place special emphasis on the chemical transformations taking place as citrate formed from acetyl-CoA and oxaloacetate is oxidized to yield CO2 and the energy of this oxidation is conserved in the reduced coenzymes NADH and FADH2.

Formation of Citrate  The first reaction of the cycle is the condensation of acetyl-CoA with oxaloacetate to form citrate, catalyzed by citrate synthase:

In this reaction the methyl carbon of the acetyl group is joined to the carbonyl group (C-2) of oxaloacetate. Citroyl-CoA is a transient intermediate. It is formed on the active site of the enzyme and rapidly undergoes hydrolysis to yield free CoA and citrate, which are then released from the active site. The hydrolysis of the high-energy thioester intermediate makes the forward reaction highly exergonic. The large, negative free-energy change associated with the citrate synthase reaction is essential to the operation of the cycle, because of the very low concentration of oxaloacetate normally present. The CoA formed in this reaction is recycled; it is ready to participate in the oxidative decarboxylation of another molecule of pyruvate by the pyruvate dehydrogenase complex to yield another molecule of acetyl-CoA for entry into the cycle.

The citrate synthase from mitochondria has been crystallized and visualized by x-ray crystallography in the presence and absence of its substrates and inhibitors. Oxaloacetate, the first substrate to bind to the enzyme, induces a large conformational change, creating a binding site for the second substrate, acetyl-CoA. When citroyl-CoA forms on the enzyme surface, another conformational change brings the side chain of a crucial Asp residue into position to cleave the thioester. This induced fit of the enzyme first to its substrate and then to its intermediate decreases the likelihood of premature and unproductive cleavage of the thioester bond of acetyl-CoA.

Formation of Isocitrate via cis-Aconitate  The enzyme aconitase (more formally, aconitate hydratase) catalyzes the reversible transformation of citrate to isocitrate, through the intermediary formation of the tricarboxylic acid cis-aconitate, which normally does not dissociate from the active site. Aconitase can promote the reversible addition of H2O to the double bond of enzyme-bound cis-aconitate in two different ways, one leading to citrate and the other to isocitrate:

Figure 15–8  The iron–sulfur center in aconitase acts in substrate binding and catalysis. Three Cys residues of the enzyme bind three iron atoms in the iron–sulfur center (yellow); the fourth iron is bound to one of the carboxyl groups of citrate (blue). : B represents a basic residue on the enzyme that helps to position the citrate in the active site.
ΔG°’ = 13.3 kJ/mol

Although the equilibrium mixture at pH 7.4 and 25 °C contains less than 10% isocitrate, in the cell the reaction is pulled to the right because isocitrate is rapidly consumed in the subsequent step of the cycle, lowering its steady-state concentration. Aconitase contains an iron–sulfur center (Fig. 15–8), which acts both in the binding of the substrate at the active site and in catalysis of the addition or removal of H2O.

Oxidation of Isocitrate to α-Ketoglutarate and CO2  In the next step isocitrate dehydrogenase catalyzes oxidative decarboxylation of isocitrate to form α-ketoglutarate:

ΔG°’ = –20.9 kJ/mol

There are two different forms of isocitrate dehydrogenase (isozymes; see Box 14–3), one requiring NAD+ as electron acceptor and the other requiring NADP+. The overall reactions catalyzed by the two isozymes are otherwise identical. The NAD-dependent enzyme is found in the mitochondrial matrix and serves in the citric acid cycle to produce α-ketoglutarate. The NADP-dependent isozyme is found in both the mitochondrial matrix and the cytosol. It may function primarily in the generation of NADPH, which is essential for reductive anabolic reactions.

Oxidation of α-Ketoglutarate to Succinyl-CoA and CO2  The next step is another oxidative decarboxylation, in which α-ketoglutarate is converted to succinyl-CoA and O2 by the action of the α-ketoglutarate dehydrogenase complex; NAD+ serves as electron acceptor:

ΔG°’ = –33.5 kJ/mol

This reaction is virtually identical to the pyruvate dehydrogenase reaction discussed above; both involve the oxidation of an α-keto acid

with loss of the carboxyl group as CO2. The energy of oxidation of α-ketoglutarate is conserved in the formation of the thioester bond of succinyl-CoA. In both structure and function the α-ketoglutarate dehydrogenase complex closely resembles the pyruvate dehydrogenase complex. It includes three enzymes, analogous to E1, E2, and E3 of the pyruvate dehydrogenase complex, as well as enzyme-bound TPP, bound lipoate, FAD, NAD, and coenzyme A. Although E1 of the α-ketoglutarate dehydrogenase complex is structurally similar to E1 of pyruvate dehydrogenase, their amino acid sequences are different; the E1 components specifically bind either α-ketoglutarate or pyruvate, conferring substrate specificity upon their respective enzyme complexes. The subunits of E3 for the two complexes are virtually identical. The E2 components of the two complexes are very similar; both have covalently bound lipoyl moieties. It is a near certainty that the proteins of these two multienzyme complexes share a common evolutionary origin.

Conversion of Succinyl-CoA to Succinate  Succinyl-CoA, like acetyl-CoA, has a strongly negative free energy of hydrolysis of its thioester bond (ΔG°’ ≈ –36 kJ/mol). In the next step of the citric acid cycle, energy released in the breakage of this bond is used to drive the synthesis of a phosphoanhydride bond in either GTP or ATP, and succinate is also formed in the process:

The enzyme that catalyzes this reversible reaction is called either succinyl-CoA synthetase or succinic thiokinase; both names indicate the participation of a nucleoside triphosphate in the reaction (Box 15–1). It was long thought that the enzyme from animal tissues used GDP exclusively, whereas that from plants and bacteria used predominantly ADP. It now appears that some animal cells contain two isozymes, one specific for ADP and the other for GDP.

This energy-conserving reaction involves an intermediate step in which the enzyme molecule itself becomes phosphorylated at a His residue in the active site (Fig. 15–9). This phosphate group, which has a high group transfer potential, is transferred to ADP (or GDP) to form ATP (or GTP). The coupled formation of ATP (or GTP) at the expense of the energy released by the oxidative decarboxylation of α-ketoglutarate is another example of a substrate-level phosphorylation, like the synthesis of ATP coupled to the oxidation of glyceraldehyde-3-phosphate in glycolysis (p. 411). These reactions are called substrate-level phosphorylations to distinguish them from oxidative or respiration-linked phosphorylation. Substrate-level phosphorylations involve soluble enzymes and chemical intermediates such as the phosphohistidine residue in succinyl-CoA synthetase, or 1,3-bisphosphoglycerate in the glycolytic pathway. Respiration-linked phosphorylation, on the other hand, involves membrane-bound enzymes and transmembrane gradients of protons (Chapter 18).

The GTP formed by succinyl-CoA synthetase may donate its terminal phosphate group to ADP to form ATP, by the reversible action of nucleoside diphosphate kinase:

Thus the net result of the activity of either isozyme of succinyl-CoA synthetase is the conservation of energy as ATP. There is no change in free energy for the nucleoside diphosphate kinase reaction; ATP and GTP are energetically equivalent.

Oxidation of Succinate to Fumarate  The succinate formed from succinyl-CoA is oxidized to fumarate by the flavoprotein succinate dehydrogenase (right). In eukaryotes, succinate dehydrogenase is tightly bound to the inner mitochondrial membrane (in prokaryotes, to the plasma membrane); it is the only enzyme of the citric acid cycle that is membrane-bound. The enzyme from beef heart mitochondria contains three different iron–sulfur clusters as well as one molecule of covalently bound FAD. Electrons pass from succinate through the FAD and iron–sulfur centers before entering the chain of electron carriers
in the mitochondrial inner membrane (or the plasma membrane of bacteria). Electron flow from succinate through these carriers to the final electron acceptor, O2, is coupled to the synthesis of two ATP molecules
per pair of electrons (respiration-linked phosphorylation; Chapter 18). Malonate, an analog of succinate, is a strong competitive inhibitor of succinate dehydrogenase and therefore blocks the citric acid cycle.

Hydration of Fumarate to Produce Malate  The reversible hydration of fumarate to L-malate is catalyzed by fumarase (fumarate hydratase):

This enzyme is highly stereospecific; it catalyzes hydration of the trans double bond of fumarate but does not act on maleate, the cis isomer of fumarate. In the reverse direction (from L-malate to fumarate), fumarase is equally stereospecific: D-malate is not a substrate.

Oxidation of Malate to Oxaloacetate  In the last reaction of the citric acid cycle, NAD-linked L-malate dehydrogenase catalyzes the oxidation of L-malate to oxaloacetate:

The equilibrium of this reaction lies far to the left under standard thermodynamic conditions. However, in intact cells oxaloacetate is continually removed by the highly exergonic citrate synthase reaction (p. 454). This keeps the concentration of oxaloacetate in the cell extremely low (< 10−6 M), pulling the malate dehydrogenase reaction toward oxaloacetate formation.

B O X  15–1
Synthases, Synthetases, Ligases, Lyases, and Kinases: A Refresher on Enzyme Nomenclature

Citrate synthase is one of many enzymes that catalyze condensation reactions, yielding a product more chemically complex than its precursors. Synthases catalyze condensation reactions in which no nucleoside triphosphate (ATP, GTP, etc.) is required as an energy source. Synthetases also catalyze condensations, but this name is reserved for enzymes that do use ATP or another nucleoside triphosphate as a source of energy for the synthetic reaction. Succinyl-CoA synthetase is such an enzyme. Ligase (from the Latin ligare, meaning “to tie together”) is the general name applied to those enzymes that catalyze condensation reactions in which two atoms are joined using the energy of ATP or another energy source. DNA ligase, for example, closes breaks in DNA molecules, using energy supplied by either ATP or NAD+; it is widely used in joining DNA pieces for genetic engineering (Chapter 28). Ligases are not to be confused with lyases, enzymes that catalyze cleavages (or, in the reverse directions, additions) in which electronic rearrangements occur. The pyruvate dehydrogenase complex, which oxidatively cleaves CO2 from pyruvate, is in this large class of enzymes. The name kinase is applied to those enzymes that transfer a phosphate group from a nucleoside triphosphate such as ATP to some acceptor molecule – a sugar (as in hexokinase and glucokinase), a protein (as in glycogen phosphorylase kinase), another nucleotide (as in nucleoside diphosphate kinase), or a metabolic intermediate such as oxaloacetate (as in PEP carboxykinase).

Unfortunately, these descriptions of enzyme types overlap, and many enzymes are commonly called by two or more names. Succinyl-CoA synthetase, for example, is also called succinate thiokinase; the enzyme is clearly both a synthetase in the citric acid cycle and a kinase when acting in the direction of succinyl-CoA synthesis. This raises another source of confusion in the naming of enzymes: an enzyme is sometimes discovered by the use of an assay of the conversion of, say, A to B and is named for that reaction, but later it is found to function in the cell primarily in converting B to A. Commonly, the first name continues to be used, although the metabolic role of the enzyme would clearly be better described by naming it for the reverse reaction. The enzyme pyruvate kinase (which acts in glycolysis) illustrates this situation (p. 413). To a beginner in biochemistry, this duplication in nomenclature can be bewildering. International committees have made heroic efforts to systematize the nomenclature of enzymes (see Table 8–3 for a brief summary of the system), but often the systematic names are so long and cumbersome that they are not frequently used in biochemical conversation.

We have tried throughout this book to use the name most commonly used by working biochemists, and to point out cases in which a given enzyme has more than one widely used name. For an authoritative treatment of systematic enzyme nomenclature, see: Webb, E. (1984) Enzyme Nomenclature, Academic Press, Inc., Orlando, FL.
Figure 15–10  Each turn of the citric acid cycle produces three NADH and one FADH2, as well as one GTP (or ATP). Two CO2 are released in oxidative decarboxylation reactions. (Here and in several following figures, all cycle reactions are shown in one direction only. Keep in mind that most of the reactions are actually reversible, as shown in Fig. 15–7.)

We have now covered one complete turn of the citric acid cycle (Fig. 15–10). An acetyl group, containing two carbon atoms, was fed into the cycle by combining with oxaloacetate. Two carbon atoms emerged from the cycle as CO2 with the oxidation of isocitrate and α-ketoglutarate, and at the end of the cycle a molecule of oxaloacetate was regenerated.

Note that the two carbon atoms appearing as CO2 are not the same two carbons that entered in the form of the acetyl group; additional turns around the cycle are required before the carbon atoms that entered as an acetyl group finally appear as CO2 (Fig. 15–7).

Although the citric acid cycle itself directly generates only one molecule of ATP per turn (in the conversion of succinyl-CoA to succinate), the four oxidation steps in the cycle provide a large flow of electrons into the respiratory chain and thus eventually lead to formation of a large number of ATP molecules during oxidative phosphorylation.

We saw in the previous chapter that the energy yield from glycolysis (in which one molecule of glucose is converted into two of pyruvate) is two ATP molecules from one of glucose. When both pyruvate molecules are completely oxidized to yield six CO2 molecules in the reactions catalyzed by the pyruvate dehydrogenase complex and the enzymes of the citric acid cycle, and the electrons are transferred to O2 via the respiratory chain, as many as 38 ATP are obtained per glucose (Table 15–2). In round numbers, this represents the conservation of 38 x 30.5 kJ/mol = 1,160 kJ/mol, or 40% of the theoretical maximum of 2,840 kJ/mol available from the complete oxidation of glucose. These calculations employ the standard free-energy changes; when corrected for the actual free energy required to form ATP within cells (Box 13–2; p. 375), the calculated efficiency of the process is even greater.

Table 15–2  The stoichiometry of coenzyme reduction and ATP formation
   in the aerobic oxidation of a molecule of glucose via glycolysis, the pyruvate
   dehydrogenase reaction, and the citric acid cycle
Reaction                                                                    Number of ATP or
reduced coenzymes
directly formed
Number of ATP
Glucose   →   glucose-6-phosphate –1 ATP –1     
Fructose-6-phosphate   →   fructose-1,6-bisphosphate –1 ATP –1     
2 Glyceraldehyde-3-phosphate   →   2 1,3-bisphosphoglycerate    2 NADH 6     
2 1,3-Bisphosphoglycerate   →   2 3-phosphoglycerate 2 ATP 2     
2 Phosphoenolpyruvate   →   2 pyruvate 2 ATP 2     
2 Pyruvate   →   2 acetyl-CoA 2 NADH 6     
2 Isocitrate   →   2 α-ketoglutarate 2 NADH 6     
2 α-Ketoglutarate   →   2 succinyl-CoA 2 NADH 6     
2 Succinyl-CoA   →   2 succinate 2 ATP (or 2 GTP) 2     
2 Succinate   →   2 fumarate 2 FADH2 4     
2 Malate   →   2 oxaloacetate 2 NADH 6     
     Total      38     

* This is calculated as 3 ATP per NADH and 2 ATP per FADH2. A negative value indicates consumption.

The historical road that led to the understanding of this cyclic metabolic pathway is instructive; it illustrates the general strategies used by biochemists since early in this century to unravel complex metabolic relationships. The citric acid cycle was first postulated as the pathway of pyruvate oxidation in animal tissues by Hans Krebs, in 1937. The idea of the cycle came to him during a study of the effect of the anions of various organic acids on the rate of oxygen consumption during pyruvate oxidation by suspensions of minced pigeon-breast muscle. This muscle, used in flight, has a very high rate of respiration and was thus especially appropriate for the study of oxidative activity. Earlier investigators, particularly Albert Szent-Györgyi, had found that certain four-carbon dicarboxylic acids known to be present in animal tissues – succinate, fumarate, malate, and oxaloacetate – stimulate the consumption of oxygen by muscle. Krebs confirmed this observation and found that they also stimulate the oxidation of pyruvate. Moreover, he found that oxidation of pyruvate by muscle is also stimulated by the six-carbon tricarboxylic acids citrate, cis-aconitate, and isocitrate, and the five-carbon α-ketoglutarate. No other naturally occurring organic acids that were tested possessed such activity. The stimulatory effect of the active acids was remarkable: the addition of even a small quantity of any one of them could promote the oxidation of many times that amount of pyruvate.

The second important observation made by Krebs was that malonate (p. 458), a close analog of succinate and a competitive inhibitor of succinate dehydrogenase, inhibits the aerobic utilization of pyruvate by muscle suspensions, regardless of which active organic acid is added. This indicated that succinate and succinate dehydrogenase must be essential components in the enzymatic reactions involved in the oxidation of pyruvate. Krebs further found that when malonate is used to inhibit the aerobic utilization of pyruvate by a suspension of muscle tissue, there is an accumulation of citrate, α-ketoglutarate, and succinate in the suspending medium. This suggested that citrate and α-ketoglutarate are normally precursors of succinate.

From these basic observations and other evidence, Krebs concluded that the active tricarboxylic and dicarboxylic acids listed above can be arranged in a logical chemical sequence. Because the incubation of pyruvate and oxaloacetate with ground muscle tissue resulted in accumulation of citrate in the medium, Krebs reasoned that this sequence functions in a circular rather than linear manner: its beginning and end are linked together. For the missing link that closes the circle he proposed the reaction

Pyruvate + oxaloacetate   →   citrate + CO2

(Notice that in this detail Krebs was originally incorrect.)

From these simple experiments and logical reasoning, Krebs postulated what he called the citric acid cycle as the main pathway for oxidation of carbohydrate in muscle. The citric acid cycle is also called the tricarboxylic acid cycle because for some years after Krebs postulated the cycle it was uncertain whether citrate or some other tricarboxylic acid such as isocitrate was the first product formed by reaction of pyruvate and oxaloacetate. In the years since its discovery, the citric acid cycle has been found to function not only in muscles but in virtually all tissues of aerobic animals and plants and in many aerobic microorganisms.

The citric acid cycle was first postulated from experiments carried out on suspensions of minced muscle tissue. Subsequently its details were worked out by study of the highly purified enzymes of the cycle. One might ask whether these enzymes really function in a cycle in intact living cells and whether the rate of the cycle is high enough to account for the overall rate of glucose oxidation in animal tissues. These questions have been studied by the use of isotopically labeled metabolites, such as pyruvate or acetate, in which the isotopes 13C or 14C are used to mark a given carbon atom in the molecule. Many stringent experiments with the isotope tracer technique have confirmed that the citric acid cycle does take place in living cells, and at a high rate.

Some of the earliest experiments with isotopes produced an unexpected result, however, which aroused considerable controversy about the pathway and mechanism of the citric acid cycle. In fact, these experiments at first seemed to show that citrate was not the first tricarboxylic acid to be formed. Box 15–2 gives some details of this episode in the scientific history of the cycle.

Figure 15–11  The noncyclic reactions (blue arrows) that provide biosynthetic precursors in anaerobically growing bacteria. These cells lack α-ketoglutarate dehydrogenase and therefore cannot carry out the complete citric acid cycle, which normally follows the direction shown with gray arrows. α-Ketoglutarate and succinyl-CoA serve as precursors in a variety of biosynthetic reactions (see Fig. 15–12).

This eight-step, cyclic process for oxidation of the simple two-carbon acetyl groups to CO2 may seem unnecessarily cumbersome and not in keeping with the principle of maximum economy in the molecular logic of living cells. The role of the citric acid cycle is not confined to the oxidation of acetate, however; this pathway is the hub of intermediary metabolism. Four- and five-carbon end products of many catabolic processes are fed into the cycle to serve as fuels. Oxaloacetate and α-ketoglutarate are, for example, produced from asparate and glutamate, respectively, when proteins in the diet are degraded. Under other circumstances, intermediates are drawn out of the cycle to be used as precursors in a variety of biosynthetic pathways.

The pathway used in modern organisms is the product of evolution, much of which occurred before the advent of aerobic organisms. It does not necessarily represent the shortest pathway from acetate to CO2, but is rather the pathway that has conferred the greatest selective advantage on its possessors throughout evolution. Early anaerobes very probably used some of the reactions of the citric acid cycle in linear biosynthetic processes. In fact, there are modern anaerobic microorganisms in which an incomplete citric acid cycle serves as a source, not of energy, but of biosynthetic precursors (Fig. 15–11). These organisms use the first three reactions of the citric acid cycle to make α-ketoglutarate, but they lack α-ketoglutarate dehydrogenase and therefore cannot carry out the complete set of citric acid cycle reactions. They do have the four enzymes that catalyze the reversible conversion of oxaloacetate into succinyl-CoA (Fig. 15–11), and these anaerobes make malate, fumarate, succinate, and succinyl-CoA from oxaloacetate in a reversal of the “normal” (oxidative) direction of flow through the cycle.

With the evolution of cyanobacteria that produced O2 from water, the earth’s atmosphere became aerobic and there was selective pressure on organisms to develop aerobic metabolism, which, as we have seen, is much more efficient than anaerobic fermentation.

B O X  15–2
Is Citric Acid the First Tricarboxylic Acid Formed in the Cycle?

When the heavy-carbon isotope 13C and the radioactive carbon isotopes 11C and 14C became available, they were very soon put to use to trace the pathway of carbon atoms through the citric acid cycle. In one such experiment, which initiated the controversy over the role of citric acid, acetate labeled in the carboxyl group (designated [1-14C]acetate) was incubated aerobically with an animal tissue preparation. Acetate is enzymatically converted into acetyl-CoA in animal tissues, and the pathway of the labeled carboxyl carbon of the acetyl group in the cycle reactions could be traced. α-Ketoglutarate was isolated from the tissue after incubation, then degraded by known chemical reactions to establish the position(s) of the isotopic carbon derived from carboxyl-labeled acetate. Condensation of unlabeled oxaloacetate with carboxyl-labeled acetate would be expected to produce citrate labeled in one of the two primary carboxyl groups (Fig. 1). Because citrate is a symmetric molecule, with no asymmetric carbon, its two terminal carboxyl groups are chemically indistinguishable. Therefore, half of the labeled citrate molecules were expected to yield α-ketoglutarate labeled in the α-carboxyl group and the other half to yield α-ketoglutarate labeled in the γ-carboxyl group; that is, the α-ketoglutarate isolated should have been a mixture of molecules labeled in both carboxyl groups.

Contrary to this expectation, the labeled α-ketoglutarate isolated from the tissue suspension contained the isotope only in the γ-carboxyl group (Fig. 1). It was concluded that citrate itself, or any other symmetric molecule, could not possibly be an intermediate in the pathway from acetate to α-ketoglutarate. Hence an asymmetric tricarboxylic acid, presumably cis-aconitate or isocitrate, had to be the first condensation product formed from acetate and oxaloacetate.

In 1948, however, Alexander Ogston pointed out that although citrate has no chiral center (see Fig. 3–9), it has the potential of reacting asymmetrically if the enzyme acting upon it has an active site that is asymmetric. He suggested that the active site of aconitase, the enzyme acting on the newly formed citrate, may have three points to which the citrate molecule must be bound and that the citrate molecule must undergo a specific three-point attachment to these binding points. As seen in Figure 2, the binding of citrate to the three points can happen in only one way, and this would account for the formation of only one type of labeled α-ketoglutarate. Organic molecules such as citrate that have no chiral center, but are potentially capable of reacting asymmetrically with an asymmetric active site, are now called prochiral molecules.

Figure 15–12  Intermediates of the citric acid cycle are drawn off as precursors in many biosynthetic pathways, yielding the products in the shaded areas. Shown in red are four anaplerotic reactions that replenish depleted intermediates of the citric acid cycle (see Table 15-3).

In aerobic organisms, the citric acid cycle is an amphibolic pathway (i.e., it serves in both catabolic and anabolic processes). It not only functions in the oxidative catabolism of carbohydrates, fatty acids, and amino acids, but also provides precursors for many biosynthetic pathways (Fig. 15–12), as in anaerobic ancestors. By the action of several important auxiliary enzymes, certain intermediates of the citric acid cycle, particularly α-ketoglutarate and oxaloacetate, can be removed from the cycle to serve as precursors of amino acids. Aspartate and glutamate have the same carbon skeletons as oxaloacetate and α-ketoglutarate, respectively, and are synthesized from them by simple transamination (Chapter 21). Through aspartate and glutamate the carbons of oxaloacetate and α-ketoglutarate are used to build other amino acids as well as purine and pyrimidine nucleotides. We will see in Chapter 19 how oxaloacetate is converted into glucose in the process of gluconeogenesis. Succinyl-CoA is a central intermediate in the synthesis of the porphyrin ring of heme groups, which serve as oxygen carriers (in hemoglobin and myoglobin) and electron carriers (in cytochromes).

Given the number of biosynthetic products derived from citric acid cycle intermediates, this cycle clearly serves a critical role apart from its function in energy-yielding metabolism.

When intermediates of the citric acid cycle are removed to serve as biosynthetic precursors, the resulting decrease in the concentration of these intermediates would be expected to slow the flux through the citric acid cycle. However, the intermediates can be replenished by anaplerotic reactions (Fig. 15–12; Table 15–3). Under normal circumstances the reactions by which the cycle intermediates are drained away and those by which they are replenished are in dynamic balance, so that the concentrations of the citric acid cycle intermediates remain almost constant.

In animal tissues one important anaplerotic reaction is the reversible carboxylation of pyruvate by CO2 to form oxaloacetate, catalyzed by pyruvate carboxylase (Table 15–3). The three other anaplerotic reactions shown in Table 15–3 also serve, in various tissues and organisms, to convert either pyruvate or phosphoenolpyruvate to oxaloacetate. When the citric acid cycle is deficient in oxaloacetate or any of the other intermediates, pyruvate is carboxylated to produce more oxaloacetate. The enzymatic addition of a carboxyl group to the pyruvate molecule requires energy, which is supplied by ATP. Because the standard free-energy change of the overall reaction is very small, we can conclude that the free energy required to attach a carboxyl group to pyruvate is about equal to the free energy available from ATP. The carboxylation of pyruvate also requires the vitamin biotin (Fig. 15–13a), which is the prosthetic group of pyruvate carboxylase.

The pyruvate carboxylase reaction is the most important anaplerotic reaction in the liver and kidney of mammals. Other anaplerotic reactions are more important in other tissues and organisms (Table 15–3).

Pyruvate carboxylase is a regulatory enzyme and is virtually inactive in the absence of acetyl-CoA, its positive allosteric modulator. Whenever acetyl-CoA, which is the fuel for the citric acid cycle, is present in excess, it stimulates the pyruvate carboxylase reaction to produce more oxaloacetate, enabling the cycle to use more acetyl-CoA in the citrate synthase reaction.

The other anaplerotic reactions are also regulated to keep the level of intermediates high enough to support the activity of the citric acid cycle. Phosphoenolpyruvate carboxylase, for example, is activated by the glycolytic intermediate fructose-1,6-bisphosphate, the level of which rises when the citric acid cycle operates too slowly to process the pyruvate generated by glycolysis.

Biotin plays a key role in many carboxylation reactions. This vitamin is a specialized carrier of one-carbon groups in their most oxidized form: CO2. (The transfer of one-carbon groups in more reduced forms is mediated by other cofactors, notably tetrahydrofolate and S-adenosylmethionine, as described in Chapter 17.) Carboxyl groups are attached to biotin at the ureido group within the biotin ring system (Fig. 15–13a).

Pyruvate carboxylase is composed of four identical subunits, each containing a molecule of biotin covalently attached through an amide linkage between its valerate side chain and the ϵ-amino group of a specific Lys residue in the enzyme active site (Fig. 15–13b); this biotinyllysine is called biocytin. Carboxylation of pyruvate proceeds in two steps; first, a carboxyl group derived from HCO3 is attached to biotin, then the carboxyl group is transferred to pyruvate to form oxaloacetate. These two steps occur at separate active sites; the long flexible arm of biocytin permits the transfer of activated carboxyl groups from the first active site to the second, much as the long lipoyllysyl arm of E2 functions in the pyruvate dehydrogenase complex.

Biotin is a vitamin required in the human diet; it is abundant in many foods and is synthesized by intestinal bacteria. Deficiency diseases are rare and are generally observed only when large quantities of raw eggs are consumed. Egg whites contain a large amount of the protein avidin (Mr 70,000), which binds to biotin very tightly and prevents its absorption in the intestine. The avidin in egg whites may be a defense mechanism, inhibiting the growth of bacteria. When eggs are cooked, avidin is denatured (and thereby inactivated) along with all other egg white proteins.

We saw in Chapter 14 that key enzymes in metabolic pathways are regulated by allosteric effectors and by covalent modification, to assure production of intermediates and products at the rates required to keep the cell in a stable steady state and to avoid wasteful overproduction of intermediates. The flow of carbon atoms from pyruvate into and through the citric acid cycle is under tight regulation at two levels: the conversion of pyruvate into acetyl-CoA, the starting material for the cycle (the pyruvate dehydrogenase complex reaction), and the entry of acetyl-CoA into the cycle (the citrate synthase reaction). Because pyruvate is not the sole source of acetyl-CoA (most cells can obtain acetyl-CoA by the oxidation of fatty acids and certain amino acids), the availability of intermediates from these other pathways is also important in the regulation of pyruvate oxidation and of the citric acid cycle. The cycle is also regulated at the isocitrate dehydrogenase and α-ketoglutarate dehydrogenase reactions.

Figure 15–14  Regulation of metabolite flow from pyruvate through the citric acid cycle. The pyruvate dehydrogenase complex is allosterically inhibited at high [ATP]/[ADP], [NADH]/[NAD+], and [acetyl-CoA]/[CoA] ratios, all of which indicate the energy-sufficient metabolic state. When these ratios decrease, allosteric activation of pyruvate oxidation results. The rate of flow through the citric acid cycle can be limited by the availability of the substrates oxaloacetate and acetyl-CoA or by the depletion of NAD+ by its conversion to NADH, which slows the three oxidation steps for which NAD+ is the cofactor. Feedback inhibition by succinyl-CoA, citrate, and ATP also slows the cycle by inhibiting early steps. In muscle tissue, Ca2+ signals contraction and stimulates energy-yielding metabolism to replace the ATP consumed by contraction.

The pyruvate dehydrogenase complex of vertebrates is regulated both allosterically and by covalent modification. The complex is strongly inhibited by ATP, as well as by acetyl-CoA and NADH, the products of

the reaction (Fig. 15–14). The allosteric inhibition of pyruvate oxidation is greatly enhanced when long-chain fatty acids are available. AMP, CoA, and NAD+, all of which accumulate when too little acetate flows into the citric acid cycle, allosterically activate the pyruvate dehydrogenase complex. Thus this enzyme activity is turned off when ample fuel is available in the form of fatty acids and acetyl-CoA and when the cell’s ATP concentration and [NADH]/[NAD+] ratio are high, and turned on when energy demands are high and greater flux of acetyl-CoA into the citric acid cycle is required.

In the pyruvate dehydrogenase complex of vertebrates, these allosteric regulation mechanisms are complemented by a second level of regulation, covalent protein modification. The enzyme complex is inhibited by the reversible phosphorylation of a specific Ser residue on one of the two subunits of E1. As noted earlier, in addition to the enzymes E1, E2, and E3, the pyruvate dehydrogenase complex contains two regulatory proteins, the sole purpose of which is to regulate the activity of the complex. A specific protein kinase phosphorylates and thereby inactivates E1, and a specific phosphoprotein phosphatase removes the phosphate group by hydrolysis, thereby activating E1. The kinase is allosterically activated by ATP: when ATP levels are high (reflecting a sufficient supply of energy), the pyruvate dehydrogenase

complex is inactivated by phosphorylation of E1. When the ATP level declines, kinase activity decreases and phosphatase action removes the phosphates from E1, activating the complex.

The pyruvate dehydrogenase complex of plants, which is found in the mitochondrial matrix and within plastids (p. 39), is strongly inhibited by NADH, which may be its primary regulator. There is also evidence for inactivation of the plant mitochondrial enzyme by reversible phosphorylation. The pyruvate dehydrogenase complex of E. coli is under allosteric regulation similar to that of the vertebrate enzyme, but the regulation by phosphorylation apparently does not occur with the bacterial enzyme.

The flow of metabolites through the citric acid cycle is under stringent, but not complex, regulation. Three factors govern the rate of flux through the cycle: substrate availability, inhibition by accumulating products, and allosteric feedback inhibition of early enzymes by later intermediates in the cycle.

There are three strongly exergonic steps in the cycle, those catalyzed by citrate synthase, isocitrate dehydrogenase, and α-ketoglutarate dehydrogenase (Fig. 15–14). Each can become the rate-limiting step under some circumstances. The availability of the substrates for citrate synthase (acetyl-CoA and oxaloacetate) varies with the metabolic circumstances and sometimes limits the rate of citrate formation. NADH, a product of the oxidation of isocitrate and α-ketoglutarate, accumulates under some conditions, and when the [NADH]/[NAD+] ratio becomes large, both dehydrogenase reactions are severely inhibited by mass action. Similarly, the malate dehydrogenase reaction is essentially at equilibrium in the cell (i.e., it is substrate limited; see Fig. 14–16), and when [NADH]/[NAD+] is large, the concentration of oxaloacetate is low, slowing the first step in the cycle. Product accumulation inhibits all three of the limiting steps of the cycle: succinyl-CoA inhibits α-ketoglutarate dehydrogenase (and also citrate synthase); citrate blocks citrate synthase; and the end product, ATP, inhibits both citrate synthase and isocitrate dehydrogenase (Fig. 15–14). The inhibition of citrate synthase by ATP is relieved by ADP, an allosteric activator of this enzyme. Calcium ions, which in vertebrate muscle are the signal for contraction and the concomitant increased demand for ATP, activate both isocitrate dehydrogenase and α-ketoglutarate dehydrogenase, as well as the pyruvate dehydrogenase complex. In short, the concentrations of substrates and intermediates of the citric acid cycle set the flux through this pathway at a rate that provides optimal concentrations of ATP and NADH.

Under normal conditions the rates of glycolysis and of the citric acid cycle are integrated so that only as much glucose is metabolized to pyruvate as is needed to supply the citric acid cycle with its fuel, the acetyl groups of acetyl-CoA. Pyruvate, lactate, and acetyl-CoA are normally maintained at steady-state concentrations. The rate of glycolysis is matched to the rate of the citric acid cycle not only by its inhibition by high levels of ATP and NADH, which are common components of both the glycolytic and respiratory stages of glucose oxidation, but also by the concentration of citrate. Citrate, the product of the first step of the citric acid cycle, serves as an important allosteric inhibitor of the phosphorylation of fructose-6-phosphate by phosphofructokinase-1 in the glycolytic pathway (p. 434).

The conversion of phosphoenolpyruvate to pyruvate (p. 413) and of pyruvate to acetyl-CoA (Fig. 15–2) are so exergonic as to be essentially irreversible. If a cell cannot convert acetate into phosphoenolpyruvate, acetate cannot serve as the starting material for the gluconeogenic pathway that leads from phosphoenolpyruvate to glucose (Chapter 19). Without this capacity, a cell or organism is unable to convert fuels that are degraded to acetate (fatty acids and certain amino acids) into carbohydrates.

As we saw in our discussion of anaplerotic reactions (Table 15–3), phosphoenolpyruvate can be synthesized from oxaloacetate in the reversible reaction catalyzed by PEP carboxykinase:

Oxaloacetate + GTP   ⇌   phosphoenolpyruvate + CO2 + GDP

In Chapter 19 we will see how phosphoenolpyruvate is converted to glucose by the gluconeogenic pathway.

Because carbon atoms from acetate molecules that enter the citric acid cycle appear eight steps later in oxaloacetate, it might appear that operation of the citric acid cycle could generate oxaloacetate from acetate, and thus generate phosphoenolpyruvate for gluconeogenesis. However, examination of the stoichiometry of the cycle reveals that there is no net conversion of acetate into oxaloacetate via the cycle; for every two carbons that enter the cycle as acetyl-CoA, two leave as CO2.

In plants, in certain invertebrates, and in some microorganisms such as E. coli and yeast, acetate can serve both as an energy-rich fuel and as a source of phosphoenolpyruvate for carbohydrate synthesis. These organisms have a pathway, the glyoxylate cycle, that allows the net conversion of acetate to oxaloacetate. In these organisms, some enzymes of the citric acid cycle operate in two modes: (1) they can function in the citric acid cycle for the oxidation of acetyl-CoA to CO2, as it occurs in most tissues, and (2) they can operate as part of a specialized modification, the glyoxylate cycle (Fig. 15–15). The glyoxylate cycle may have evolved before, and given rise to, the citric acid cycle. The overall reaction equation of the glyoxylate cycle, which may also be regarded as an anaplerotic pathway, is

2 Acetyl-CoA + NAD+ + 2H2O   →   succinate + 2CoA + NADH + H+
Figure 15–15  The glyoxylate cycle and its relationship to the citric acid cycle. The orange reaction arrows represent the glyoxylate cycle, and the blue arrows, the citric acid cycle. Notice that the glyoxylate cycle bypasses the two decarboxylation steps of the citric acid cycle, and that two molecules of acetyl-CoA enter the glyoxylate cycle during each turn, but only one enters the citric acid cycle. The glyoxylate cycle was elucidated by Hans Kornberg and Neil Madsen in the laboratory of Hans Krebs.
In the glyoxylate cycle, acetyl-CoA condenses with oxaloacetate to form citrate exactly as in the citric acid cycle. The breakdown of isocitrate does not occur via the isocitrate dehydrogenase reaction, however, but through a cleavage catalyzed by the enzyme isocitrate lyase, to form succinate and glyoxylate. The glyoxylate then condenses with acetyl-CoA to yield malate in a reaction catalyzed by malate synthase. The malate is subsequently oxidized to oxaloacetate, which can condense with another molecule of acetyl-CoA to start another turn of the cycle (Fig. 15–15). In each turn of the glyoxylate cycle, two molecules of acetyl-CoA enter and there is a net synthesis of one molecule of succinate, available for biosynthetic purposes. The succinate may be converted through fumarate and malate into oxaloacetate, which can then be converted into phosphoenolpyruvate by the PEP carboxykinase reaction described above. Phosphoenolpyruvate can then serve as a precursor of glucose in gluconeogenesis.
In plants, the enzymes of the glyoxylate cycle are sequestered in membrane-bounded organelles called glyoxysomes (Fig. 15–16); those enzymes common to the citric acid and glyoxylate cycles have two isozymes, one specific to mitochondria, the other to glyoxysomes. Glyoxysomes are not present in all plant tissues at all times. They develop in lipid-rich seeds during germination, before the developing plants acquire the ability to make glucose by photosynthesis. In addition to glyoxylate cycle enzymes, glyoxysomes also contain all of the enzymes needed for the degradation of fatty acids stored in seed oils (Chapter 16). Acetyl-CoA formed from lipids is converted into malate via the glyoxylate cycle, and the malate serves as a source of oxaloacetate (through the malate dehydrogenase reaction) for gluconeogenesis. Germinating plants are therefore able to convert the carbon of seed lipids into glucose.

Vertebrate animals do not have the enzymes specific to the glyoxylate cycle (isocitrate lyase and malate synthase) and therefore cannot bring about the net synthesis of glucose from lipids.

Figure 15–17  The reactions of the glyoxylate cycle (in glyoxysomes) proceed simultaneously with, and mesh with, those of the citric acid cycle (in mitochondria), as intermediates pass through the cytosol between these compartments. The reactions involved in the oxidation of fatty acids to acetyl-CoA and the conversion of oxaloacetate to aspartate will be discussed in Chapters 16 and 21, respectively.

In germinating plant seeds, the enzymatic transformations of dicarboxylic and tricarboxylic acids occur in three intracellular compartments: mitochondria, glyoxysomes, and the cytosol. There is a continuous interchange of intermediates among these compartments (Fig. 15–17).

Aspartate carries the carbon skeleton of oxaloacetate from the citric acid cycle (in mitochondria) to the glyoxysome, where it condenses with acetyl-CoA derived from fatty acid breakdown. The citrate thus formed is converted to isocitrate by aconitase, then split into glyoxylate and succinate by isocitrate lyase. The succinate returns to the mitochondrion, where it reenters the citric acid cycle and is transformed into oxaloacetate, which can again be exported (via aspartate) to the glyoxysome. The glyoxylate formed within the glyoxysome combines with acetyl-CoA to yield malate, which enters the cytosol and is oxidized (by cytosolic malate dehydrogenase) to oxaloacetate, the precursor of glucose via gluconeogenesis. Four distinct pathways participate in these conversions: fatty acid breakdown to acetyl-CoA (in glyoxysomes),

the glyoxylate cycle (in glyoxysomes), the citric acid cycle (in mitochondria), and gluconeogenesis (in the cytosol).
Figure 15–18  Regulation of isocitrate dehydrogenase activity determines the partitioning of isocitrate between the glyoxylate cycle and the citric acid cycle. When isocitrate dehydrogenase is inactivated by phosphorylation (by a specific protein kinase), isocitrate is directed into biosynthetic reactions via the glyoxylate cycle; when the enzyme is activated by dephosphorylation (by a specific phosphatase), isocitrate enters the citric acid cycle, and ATP production results.
The sharing of common intermediates requires that these pathways be regulated and coordinated. Isocitrate is a crucial intermediate, standing at the branch point between the glyoxylate and citric acid cycles (Fig. 15–18). Isocitrate dehydrogenase is regulated by covalent modification: a specific protein kinase phosphorylates and thereby inactivates the dehydrogenase. Inactivation of isocitrate dehydrogenase shunts isocitrate to the glyoxylate cycle, where it begins the synthetic route toward glucose. A phosphoprotein phosphatase removes the phosphate group from isocitrate dehydrogenase, reactivating the enzyme and sending more isocitrate through the energy-yielding citric acid cycle. The regulatory protein kinase and phosphoprotein phosphatase are separate enzymatic activities, but both reside in the same polypeptide.

Some bacteria, including E. coli, have the full complement of enzymes for the glyoxylate and citric acid cycles in the cytosol. E. coli can therefore grow with acetate as its sole source of carbon and energy. The phosphatase activity that causes activation of isocitrate dehydrogenase is stimulated by intermediates of the citric acid cycle and of glycolysis, and by indicators of reduced cellular energy supply (Table 15–4; Fig. 15–18). The same metabolites inhibit the protein kinase activity of the bifunctional enzyme. Thus, the accumulation of intermediates of the central energy-yielding pathways, or energy depletion, results in the activation of isocitrate dehydrogenase. When the concentration of these regulators falls, signaling enough flux through the energy-yielding citric acid cycle, isocitrate dehydrogenase is inactivated by the protein kinase.

Table 15–4  Allosteric effectors of the regulatory kinase/phosphatase
protein of isocitrate dehydrogenase
Citric acid
cycle intermediates           
Glycolytic intermediates Cofactors that indicate
energy depletion
Citrate Phosphoenolpyruvate* AMP*
Isocitrate* Pyruvate* ADP*
α-Ketoglutarate* 3-Phosphoglycerate* NADP+
Oxaloacetate* Fructose-6-phosphate

All compounds shown inhibit the kinase activity. Compounds with * stimulute the phosphatase
activity. The overall result is activation of isocitrate dehydrogenase and thus of the citric acid

The same intermediates of the glycolytic and citric acid cycles that lead to activation of isocitrate dehydrogenase are allosteric inhibitors of isocitrate lyase. When energy-yielding metabolism is sufficiently fast to keep the concentrations of intermediates of glycolysis and the citric acid cycle low, isocitrate dehydrogenase is inactivated, the inhibition of isocitrate lyase is relieved, and isocitrate flows into the glyoxylate pathway, where it is used in the biosynthesis of carbohydrates, amino acids, and other cellular components.
Chapter 16
Oxidation of Fatty Acids

The oxidation of long-chain fatty acids to acetyl-CoA is a central energy-yielding pathway in animals, many protists, and some bacteria. The electrons removed during fatty acid oxidation pass through the mitochondrial respiratory chain, driving ATP synthesis, and the acetyl-CoA produced from the fatty acids may be completely oxidized to CO2 via the citric acid cycle, resulting in further energy conservation. In some organisms, acetyl-CoA produced by fatty acid oxidation has alternative fates. In vertebrate animals, acetyl-CoA may be converted in the liver into ketone bodies – water-soluble fuels exported to the brain and other tissues when glucose is not available. In higher plants, acetyl-CoA from fatty acid oxidation serves primarily as a biosynthetic precursor, and only secondarily as fuel. Although the biological role of fatty acid oxidation differs from organism to organism, the mechanism is essentially the same. This chapter is centered on the four-step process, called β oxidation, by which fatty acids are converted into acetyl-CoA.

In Chapter 9 we described the properties of triacylglycerols (also called triglycerides or neutral fats) that make them especially suitable as storage fuels. The long alkyl chains of their constituent fatty acids are essentially hydrocarbons, highly reduced structures with an energy of complete oxidation (~38 kJ/g) more than twice that for the same weight of carbohydrate or protein. Because of their hydrophobicity and extreme insolubility in water, triacylglycerols are segregated into lipid droplets, which do not raise the osmolarity of the cytosol and, unlike polysaccharides, do not contain extra weight as water of solvation. The relative chemical inertness of triacylglycerols allows their intracellular storage in large quantity without the risk of undesired chemical reactions with other cellular constituents.

The same properties that make triacylglycerols good storage compounds present problems in their role as fuels. Because of their insolubility in water, ingested triacylglycerols must be emulsified before they can be digested by water-soluble enzymes in the intestine, and triacylglycerols absorbed in the intestine or mobilized from storage tissues must be carried in the blood by proteins that counteract their insolubility. The relative stability of the C–C bonds in a fatty acid is overcome by activation of the carboxyl group at C-1 by attachment to coenzyme A, which allows stepwise oxidation of the fatty acyl group at the C-3 position. This latter carbon atom is also called the beta (β) carbon in common nomenclature, from which the oxidation of fatty acids gets its common name: β oxidation.

We begin this chapter with a brief discussion of the sources of fatty acids and the routes by which they are carried to the site of their oxidation, with special emphasis on the case of vertebrate animals. The chemical steps of fatty acid oxidation in mitochondria are then described. Three stages in this process can be distinguished: the oxidation of long-chain fatty acids to two-carbon fragments, in the form of acetyl-CoA; the oxidation of acetyl-CoA to CO2 via the citric acid cycle (Chapter 15); and the transfer of electrons from reduced electron carriers to the mitochondrial respiratory chain (Chapter 18). Our emphasis in this chapter is on the first of these stages. We consider the simple case in which a fully saturated fatty acid with an even number of carbon atoms is degraded to acetyl-CoA, then we look briefly at the extra transformations necessary for the degradation of unsaturated fatty acids and of fatty acids with an odd number of carbons. Finally, we discuss variations on the β-oxidation theme that occur in specialized organelles – peroxisomes and glyoxysomes. The chapter concludes with the description of an alternative fate for the acetyl-CoA formed by β oxidation in vertebrates: the production of ketone bodies in the liver.

Cells that derive energy from the oxidation of fatty acids may obtain those fatty acids from three sources: fats in the diet, fats stored in cells as lipid droplets, and (in animals) fats newly synthesized in one organ for export to another. Some organisms use all three sources under various circumstances, whereas others obtain fatty acids from only one or two of these sources. Vertebrates, for example, obtain fats in the diet, mobilize fats stored in specialized tissue (adipose tissue), and convert excess dietary carbohydrates to fats in the liver for export to other tissues. On the average, 40% or more of the daily energy requirement of humans in highly industrialized countries is supplied by dietary triacylglycerols (although most nutritional guidelines recommend that no more than 30% of the daily caloric intake be from fats). Triacylglycerols provide more than half the energy requirements of some organs, particularly the liver, heart, and resting skeletal muscle. Stored triacylglycerols are virtually the sole source of energy in hibernating animals and migrating birds. Protists obtain fats by consuming organisms lower in the food chain, and some also store fats in cytosolic lipid droplets. Higher plants mobilize fats stored in seeds during the process of germination, but do not otherwise depend on fats for energy.

Figure 16–1  Uptake of dietary lipid in the intestine of a vertebrate animal, and delivery of fatty acids to muscle and adipose tissues. The eight steps are discussed in the text.
Before ingested triacylglycerols can be absorbed through the intestinal wall, they must be converted from insoluble macroscopic fat particles into finely dispersed microscopic micelles. Bile salts such as taurocholic acid are synthesized from cholesterol in the liver, stored in the gallbladder, and released into the small intestine after ingestion of a fatty meal. These amphipathic compounds act as biological detergents, converting dietary fats into mixed micelles of bile salts and triacylglycerols (Fig. 16–1, step ). Micelle formation enormously increases the fraction of lipid molecules accessible to the action of
water-soluble lipases in the intestine, and lipase action converts triacylglycerols into monoacylglycerols (monoglycerides) and diacylglycerols (diglycerides), free fatty acids, and glycerol (step ). These products of lipase action diffuse into the epithelial cells lining the intestinal surface (the intestinal mucosa) (step ), where they are reconverted to triacylglycerols and packaged with dietary cholesterol and specific proteins into lipoprotein aggregates called chylomicrons (Fig. 16–2; see also Fig. 16–1, step ).

Apolipoproteins are lipid-binding proteins in the blood, responsible for the transport of triacylglycerols, phospholipids, cholesterol, and cholesteryl esters between organs. Apolipoproteins (“apo” designates the protein in its lipid-free form) combine with various lipids to form several classes of lipoprotein particles, spherical aggregates with hydrophobic lipids at the core and hydrophilic protein side chains and lipid head groups at the surface. Various combinations of lipid and protein produce particles of different densities, ranging from chylomicrons and very low-density lipoproteins (VLDL) to very high-density

lipoproteins (VHDL), which may be separated by ultracentrifugation. The structures and roles of these lipoprotein particles in lipid transport are detailed in Chapter 20.

The protein moieties of lipoproteins act as points of specific recognition by receptors on cell surfaces. In lipid uptake from the intestine (Fig. 16–1), chylomicrons, which contain apoprotein C-II (apoC-II), move from the intestinal mucosa into the lymphatic system, from which they enter the blood and are carried to muscle and adipose tissue (step ). In the capillaries of these tissues, the extracellular enzyme lipoprotein lipase is activated by apoC-II. This enzyme hydrolyzes triacylglycerols to fatty acids and glycerol (step ), which are taken up by cells in the target tissues (step ). In muscle, the fatty acids are oxidized for energy; in adipose tissue, they are reesterified for storage as triacylglycerols (step ).

Figure 16–2  Molecular structure of a chylomicron. The surface is covered with a layer of phospholipids, with head groups facing the aqueous phase. Triacylglycerols sequestered in the interior make up more than 80% of the mass. Several apoproteins that protrude from the surface act as signals in the uptake and metabolism of chylomicron contents. The diameter of chylomicrons ranges from about 100 to about 500 nm.

The remnants of chylomicrons, depleted of most of their triacylglycerols but still containing cholesterol and the apoproteins apoE and apoB-48, travel in the blood to the liver, where they are taken up by endocytosis, triggered by their apoproteins. Triacylglycerols that enter the liver by this route may be oxidized to provide energy or to provide precursors for the synthesis of ketone bodies, as described later in this chapter. When the diet contains more fatty acids than are needed immediately for fuel or as precursors, they are converted into triacylglycerols in the liver, and the triacylglycerols are packaged with specific apolipoproteins into VLDLs. VLDLs are transported in the blood from the liver to adipose tissues, where the triacylglycerols are removed and stored in lipid droplets within adipocytes.

Figure 16–3  Mobilization of triacylglycerols stored in adipose tissue. Low levels of glucose in the blood trigger the mobilization of triacylglycerols through the action of epinephrine and glucagon on the adipocyte adenylate cyclase. The subsequent steps in mobilization are described in the text.

When hormones signal the need for metabolic energy, triacylglycerols stored in adipose tissue are mobilized (brought out of storage) and transported to those tissues (skeletal muscle, heart, and renal cortex) in which fatty acids can be oxidized for energy production. The hormones epinephrine and glucagon, secreted in response to low blood glucose levels, activate adenylate cyclase in the adipocyte plasma membrane (Fig. 16–3), raising the intracellular concentration of cAMP (see Fig. 14–18). A cAMP-dependent protein kinase, in turn, phosphorylates and thereby activates hormone-sensitive triacylglycerol lipase, which catalyzes hydrolysis of the ester linkages of triacylglycerols. The fatty acids thus released diffuse from the adipocyte into the blood, where they bind to the blood protein serum albumin. This protein (Mr 62,000), which constitutes about half of the total serum protein, binds as many as 10 fatty acids per protein monomer by noncovalent interactions. Bound to this soluble protein, the otherwise insoluble fatty acids are carried to tissues such as skeletal muscle, heart, and renal cortex. Here, fatty acids dissociate from albumin and diffuse into the cytosol of the cells in which they will serve as fuel.

About 95% of the biologically available energy of triacylglycerols resides in their three long-chain fatty acids; only 5% is contributed by the glycerol moiety. The glycerol released by lipase action is phosphorylated by glycerol kinase (Fig. 16–4), and the resulting glycerol-3-phosphate is oxidized to dihydroxyacetone phosphate. The glycolytic enzyme triose phosphate isomerase converts this compound to glyceraldehyde-3-phosphate, which is oxidized via glycolysis.

Figure 16–4  Pathway by which glycerol derived from triacylglycerols enters glycolysis.
Figure 16–5  The reactions catalyzed by acyl-CoA synthetase and inorganic pyrophosphatase. Fatty acid activation by the formation of the fatty acyl–CoA derivative occurs in two steps. First, the carboxylate ion displaces the outer two (β and γ) phosphates of ATP to form a fatty acyl–adenylate, the mixed anhydride of a carboxylic acid and a phosphoric acid. The other product is PPi, an excellent leaving group that is immediately hydrolyzed to two Pi, pulling the reaction in the forward direction. Coenzyme A carries out nucleophilic attack on the mixed anhydride, displacing AMP and forming the thioester fatty acyl–CoA. The overall reaction is highly exergonic.

The enzymes of fatty acid oxidation in animal cells are located in the mitochondrial matrix, as demonstrated in 1948 by Eugene P. Kennedy and Albert Lehninger. The free fatty acids that enter the cytosol from the blood cannot pass directly through the mitochondrial membranes, but must first undergo a series of three enzymatic reactions. The first is catalyzed by a family of isozymes present in the outer mitochondrial membrane, acyl-CoA synthetases, which promote the general reaction

Fatty acid + CoA + ATP   ⇌   fatty acyl-CoA + AMP + PPi
The different acyl-CoA synthetase isozymes act on fatty acids of short, intermediate, and long carbon chains, respectively. Acyl-CoA synthetase catalyzes the formation of a thioester linkage between the fatty acid carboxyl group and the thiol group of coenzyme A to yield a fatty acyl–CoA; simultaneously, ATP undergoes cleavage to AMP and PPi. Recall our description of this reaction in Chapter 13 to illustrate how the free energy released by cleavage of phosphoric acid anhydride bonds in ATP could be coupled to the formation of a high-energy compound (see Fig. 13–10). The reaction occurs in two steps, and involves a fatty acyl–adenylate intermediate (Fig. 16–5).

Fatty acyl–CoAs, like acetyl-CoA, are high-energy compounds; their hydrolysis to free fatty acid and CoA has a large, negative standard free-energy change (ΔG°’ ≈ –31 kJ/mol). The formation of fatty acyl–CoAs is made more favorable by the hydrolysis of two high-energy bonds in ATP; the pyrophosphate formed in the activation reaction is immediately hydrolyzed by a second enzyme, inorganic pyrophosphatase (Fig. 16–5), which pulls the preceding activation reaction in the direction of the formation of fatty acyl–CoA. The overall reaction is

Fatty acid + CoA + ATP   →   fatty acyl–CoA + AMP + 2Pi       ΔG°’1 = –32.5 kJ/mol
Fatty acyl–CoA esters formed in the outer mitochondrial membrane do not cross the inner mitochondrial membrane intact. Instead, the fatty acyl group is transiently attached to the hydroxyl group of carnitine and the fatty acyl–carnitine is carried across the inner mitochondrial membrane by a specific transporter (Fig. 16–6). In this second enzymatic reaction required for fatty acid movement into mitochondria, carnitine acyltransferase I, present on the outer face of the inner membrane, catalyzes transesterification of the fatty acyl group from coenzyme A to carnitine. The fatty acyl–carnitine ester crosses the inner mitochondrial membrane into the matrix by facilitated diffusion through the acyl-carnitine/carnitine transporter.
Figure 16–6  Fatty acid entry into mitochondria via the acyl-carnitine/carnitine transporter. After its formation at the outer surface of the inner mitochondrial membrane, fatty acyl-carnitine moves into the matrix by facilitated diffusion through the transporter. In the matrix, the acyl group is transferred back to CoA, freeing carnitine to return to the intermembrane space via the same transporter. The acyltransferase I and II enzymes are bound to the outer and inner surfaces, respectively, of the mitochondrial inner membrane. This entry process is the rate-limiting step for oxidation of fatty acids in mitochondria, as discussed later in this chapter.
In the third and final step of the entry process, the fatty acyl group is enzymatically transferred from carnitine to intramitochondrial coenzyme A by carnitine acyltransferase II. This isozyme is located on the inner face of the inner mitochondrial membrane, where it regenerates fatty acyl–CoA and releases it, along with free carnitine, into the matrix (Fig. 16–6). Carnitine reenters the space between the inner and outer mitochondrial membranes via the acyl-carnitine/carnitine transporter.

This three-step process for transferring fatty acids into the mitochondrion has the effect of separating the cytosolic and mitochondrial pools of coenzyxne A, which have different functions. The mitochondrial pool of coenzyme A is largely used in oxidative degradation of pyruvate, fatty acids, and some amino acids, whereas the cytosolic pool of coenzyme A is used in the biosynthesis of fatty acids (Chapter 20).

Once inside the mitochondrion, the fatty acyl–CoA is ready for the oxidation of its fatty acid component by a set of enzymes in the mitochondrial matrix.
Figure 16–7  Stages of fatty acid oxidation. Stage 1: A long-chain fatty acid is oxidized to yield acetyl residues in the form of acetyl-CoA. Stage 2: The acetyl residues are oxidized to CO2 via the citric acid cycle. Stage 3: Electrons derived from the oxidations of stages 1 and 2 are passed to O2 via the mitochondrial respiratory chain, providing the energy for ATP synthesis by oxidative phosphorylation.

Mitochondrial oxidation of fatty acids takes place in three stages (Fig. 16–7). In the first stage – β oxidation – the fatty acids undergo oxidative removal of successive two-carbon units in the form of acetyl-CoA, starting from the carboxyl end of the fatty acyl chain. For example, the

16-carbon fatty acid palmitic acid (palmitate at pH 7) undergoes seven passes through this oxidative sequence, in each pass losing two carbons as acetyl-CoA. At the end of seven cycles the last two carbons of palmitate (originally C-15 and C-16) are left as acetyl-CoA. The overall result is the conversion of the 16-carbon chain of palmitate to eight two-carbon acetyl-CoA molecules. Formation of each molecule of acetyl-CoA requires removal of four hydrogen atoms (two pairs of electrons and four H+) from the fatty acyl moiety by the action of dehydrogenases.

In the second stage of fatty acid oxidation the acetyl residues of acetyl-CoA are oxidized to CO2 via the citric acid cycle, which also takes place in the mitochondrial matrix. Acetyl-CoA derived from fatty acid oxidation thus enters a final common pathway of oxidation along with acetyl-CoA derived from glucose via glycolysis and pyruvate oxidation (see Fig. 15–1).

The first two stages of fatty acid oxidation produce the reduced electron carriers NADH and FADH2, which in the third stage donate electrons to the mitochondrial respiratory chain, through which the electrons are carried to oxygen (Fig. 16–7). Coupled to this flow of electrons is the phosphorylation of ADP to ATP, to be described in Chapter 18. Thus energy released by fatty acid oxidation is conserved as ATP.

We will now look in more detail at the first stage of fatty acid oxidation, for the simple case of a saturated chain with an even number of carbons, and for the slightly more complicated cases of unsaturated and odd-number chains. We then consider the regulation of fatty acid oxidation, and the β-oxidative processes occurring in organelles other than mitochondria.

Figure 16–8  The fatty acid oxidation (β-oxidation) pathway. (a) In each pass through this sequence, one acetyl residue (shaded in red) is removed in the form of acetyl-CoA from the carboxyl end of palmitate (C16), which enters as palmitoyl-CoA. (b) Six more passes through the pathway yield seven more molecules of acetyl-CoA, the seventh arising from the last two carbon atoms of the 16-carbon chain. Eight molecules of acetyl-CoA are formed in all.
Figure 16–9  Electrons removed from fatty acids during β oxidation pass into the mitochondrial respiratory chain and eventually to O2. The structures I through IV are enzyme complexes that catalyze portions of the electron transfer to oxygen. Fatty acyl–CoA dehydrogenase feeds electrons into an electron-transferring flavoprotein (ETFP) containing an iron–sulfur center, which in turn reduces a lipid-soluble electron carrier, ubiquinone (UQ, or coenzyme Q). β-Hydroxyacyl-CoA dehydrogenase transfers electrons to NAD+, and the resulting NADH is reoxidized by NADH dehydrogenase (Complex I of the respiratory chain). Propionate produced from odd-chain fatty acids is converted to succinate. Succinate dehydrogenase, which acts in the citric acid cycle (p. 457), feeds electrons into the respiratory chain at Complex II. Cytochrome c (cyt c) is a soluble electron carrier that transfers electrons between Complexes III and IV. All of these transfers are described in detail in Chapter 18.

Four enzyme-catalyzed reactions are involved in the first stage of fatty acid oxidation (Fig. 16–8a). First, dehydrogenation produces a double bond between the α and β carbon atoms (C-2 and C-3), yielding a trans2-enoyl-CoA. The symbol Δ2 designates the position of the double bond. (It may be helpful to review fatty acid nomenclature, described on p. 240.) The new double bond has the trans configuration; recall that naturally occurring unsaturated fatty acids normally have their double bonds in the cis configuration. We shall consider the significance of this difference later.

The enzyme responsible for this first step, acyl-CoA dehydrogenase, includes FAD as a prosthetic group. The electrons removed from the fatty acyl–CoA are transferred to the FAD, and the reduced form of the dehydrogenase then immediately donates its electrons to an electron carrier, the electron-transferring flavoprotein (ETFP). ETFP, an integral protein of the inner mitochondrial membrane, is one of the electron carriers of the mitochondrial respiratory chain (Fig. 16–9). The transfer of a pair of electrons from the FADH2 of acyl-CoA dehydrogenase to O2 via the respiratory chain provides the energy for the synthesis of two ATP molecules.

The oxidation catalyzed by acyl-CoA dehydrogenase is analogous to succinate dehydrogenation in the citric acid cycle (p. 457); in both reactions the enzyme is bound to the inner membrane, a double bond is introduced into a carboxylic acid between the α and β carbons, FAD is the electron acceptor, and electrons from the reaction ultimately enter the respiratory chain and are carried to O2 with the concomitant synthesis of two ATP molecules per electron pair.

In the second step of the fatty acid oxidation cycle (Fig. 16–8a), water is added to the double bond of the trans2-enoyl-CoA to form the L stereoisomer of β-hydroxyacyl-CoA (also designated β-hydroxyacyl-CoA). This reaction, catalyzed by enoyl-CoA hydratase, is formally analogous to the fumarase reaction in the citric acid cycle, in which H2O adds across an α–β double bond (p. 458).

In the third step, the L-β-hydroxyacyl-CoA is dehydrogenated to form β-ketoacyl-CoA by the action of β-hydroxyacyl-CoA dehydrogenase (Fig. 16–8a); NAD+ is the electron acceptor. This enzyme is absolutely specific for the L stereoisomer. The NADH formed in this reaction donates its electrons to NADH dehydrogenase (Complex I), an electron carrier of the respiratory chain (Fig. 16–9). Three ATP molecules are generated from ADP per pair of electrons passing from NADH to O2 via the respiratory chain. The reaction catalyzed by β-hydroxyacyl-CoA dehydrogenase is closely analogous to the malate dehydrogenase reaction of the citric acid cycle (p. 459).

The fourth and last step of the fatty acid oxidation cycle is catalyzed by acyl-CoA acetyltransferase (more commonly called thiolase), which promotes reaction of β-ketoacyl-CoA with a molecule of free coenzyme A to split off the carboxyl-terminal two-carbon fragment of the original fatty acid as acetyl-CoA. The other product is the coenzyme A thioester of the original fatty acid, now shortened by two carbon atoms (Fig. 16–8a). This reaction is called thiolysis, by analogy with the process of hydrolysis, because the β-ketoacyl-CoA is cleaved by reaction with the thiol group of coenzyme A.

The carbon–carbon single bond that connects methylene (–CH2–) groups in fatty acids is relatively stable. The β-oxidation sequence represents an elegant solution to the problem of breaking these bonds. The first three reactions of β oxidation have the effect of creating a much less stable C–C bond, in which one of the carbon atoms (the a carbon, C-2) is bonded to two carbonyl carbons. The ketone function on the β carbon (C-3) makes it a good point for nucleophilic attack by –SH of coenzyme A, catalyzed by thiolase. The acidity of the a carbon makes the terminal –CH2–CO–S-CoA a good leaving group, facilitating breakage of the α–β bond.

In one pass through the fatty acid oxidation sequence, one molecule of acetyl-CoA, two pairs of electrons, and four H+ ions are removed from the long-chain fatty acyl-CoA, to shorten it by two carbon atoms. The equation for one pass, beginning with the coenzyme A ester of our example, palmitate, is

Palmitoyl-CoA + CoA + FAD + NAD+ + H2O   →   myristoyl-CoA + acetyl-CoA + FADH2 + NADH + H+   (16–2)

Following removal of one acetyl-CoA unit from palmitoyl-CoA, the coenzyme A thioester of the shortened fatty acid remains, in this case the 14-carbon myristate. The myristoyl-CoA can now enter the β-oxidation sequence and go through another set of four reactions, exactly analogous to the first, to yield a second molecule of acetyl-CoA and lauroylCoA, the coenzyme A thioester of the 12-carbon laurate. Altogether, seven passes through the β-oxidation sequence are required to oxidize one molecule of palmitoyl-CoA to eight molecules of acetyl-CoA (Fig. 16–8b). The overall equation is

Palmitoyl-CoA + 7CoA + 7FAD + 7NAD+ + 7H2O   →   8 acetyl-CoA + 7FADH2 + 7NADH + 7H+       (16–3)

Each molecule of FADH2 formed during oxidation of the fatty acid donates a pair of electrons to ETFP of the respiratory chain (Fig. 16–9); two molecules of ATP are generated during the ensuing transfer of the electron pair to O2 and the coupled oxidative phosphorylations. Similarly, each molecule of NADH formed delivers a pair of electrons to the mitochondrial NADH dehydrogenase; the subsequent transfer of each pair of electrons to O2 results in formation of three molecules of ATP. Thus five molecules of ATP are formed for each two-carbon unit removed in one pass through the sequence as it occurs in animal tissues, such as the liver or heart. Note that water is also produced in this process. Condensation of ADP and Pi releases one H2O for each ATP formed, and transfer of electrons from NADH or FADH2 to O2 yields one H2O per electron pair. Reduction of O2 by NADH also consumes one H+ per NADH: NADH + H+ + ½O2  →  NAD+ + H2O. In hibernating animals, fatty acid oxidation provides metabolic energy, heat, and water – all essential for survival of an animal that neither eats nor drinks for long periods (Box 16–1).

The overall equation for the oxidation of palmitoyl-CoA to eight molecules of acetyl-CoA, including the electron transfers and oxidative phosphorylation, is

Palmitoyl-CoA + 7CoA + 7O2 + 35Pi + 35ADP   →   8 acetyl-CoA + 35ATP + 42H2O             (16–4)
B O X  16–1
Fat Bears Carry On β Oxidation in Their Sleep

Many animals depend on fat stores for energy during hibernation or dormancy, during migratory periods, and in other situations involving radical metabolic adjustments (as in the case of the camel, which can obtain its water supply from the oxidation of fat).

One of the most pronounced adjustments of fat metabolism occurs in the hibernation of the grizzly bear (Fig. 1). Bears go into a continuous state of dormancy for periods as long as seven months without arousal. Unlike most other hibernating species, the bear maintains its body temperature between 32 and 35 °C, nearly the normal level. Although the bear in this state expends about 6,000 kcal/day (25,000 kJ/day), it does not eat, drink, urinate, or defecate for months at a time. When accidentally aroused, the bear is almost immediately alert and ready to defend itself.

Figure 1  A grizzly bear prepares its hibernation nest, near the McNeil River in Canada.

Experimental studies have shown that the bear uses body fat as its sole fuel during hibernation. The oxidation of fat yields sufficient energy for maintaining body temperature, for active synthesis of amino acids and proteins, and for other energy-requiring activities, such as membrane transport. Fat oxidation also releases large amounts of water (p. 488), which replenishes water loss during breathing. In addition, degradation of triacylglycerols yields glycerol, which, following its enzymatic phosphorylation to glycerol-3-phosphate and oxidation to dihydroxyacetone phosphate, is converted into blood glucose. Urea formed during the degradation of amino acids is reabsorbed and recycled by the bear, the amino groups being used to make new amino acids for maintaining body proteins.

Bears store an enormous amount of body fat in preparation for their long hibernation periods. Normally, an adult grizzly bear consumes about 9,000 kcal/day during the late spring and summer. But as winter approaches bears will feed 20 hours a day and consume up to 20,000 kcal, in response to seasonal changes in hormone secretion. Large amounts of body triacylglycerols are formed from the huge amounts of carbohydrate consumed during the fattening-up period. Other hibernating species, including the tiny dormouse, also accumulate large amounts of body fat. The camel, although not a hibernator, can synthesize and store triacylglycerols in large amounts in its hump, a metabolic source of both energy and water under desert conditions.

The acetyl-CoA produced from the oxidation of fatty acids can be oxidized to CO2 and H2O by the citric acid cycle. The following equation represents the balance sheet for the second stage in the oxidation of our example, palmitoyl-CoA, together with the coupled phosphorylations of the third stage:

8 Acetyl-CoA + 16O2 + 96Pi + 96ADP   →   8CoA + 96ATP + 104H2O + 16CO2
Combining Equations 16–4 and 16–5, we obtain the overall equation for the complete oxidation of palmitoyl-CoA to carbon dioxide and water:

Palmitoyl-CoA + 23O2 + 131Pi + 131ADP   →   CoA + 131ATP + 16CO2 + 146H2O

Because the activation of palmitate to palmitoyl-CoA consumes two ATP equivalents (p. 484), the net gain per molecule of palmitate is 129 ATP. Table 16–1 summarizes the yields of NADH, FADH2, and ATP in the successive steps of fatty acid oxidation. The standard free-energy change for the oxidation of palmitate to CO2 + H2O is about 9,800 kJ/ mol. Under standard conditions, 30.5 × 129 = 3,940 kJ/mol (about 40% of the theoretical maximum) is recovered as the phosphate bond energy of ATP. However, when the free-energy changes are calculated from actual concentrations of reactants and products under intracellular conditions (see Box 13–2), the free-energy recovery is over 80%; the energy conservation is remarkably efficient.

Table 16–1  Yield of ATP during oxidation of one molecule of
   palmitoyl-CoA to CO2 and H2O
Enzyme catalyzing oxidation step    Number of NADH
or FADH2 formed 
Number of ATP
ultimately formed
Acyl-CoA dehydrogenase 7 FADH2              14     
β-Hydroxyacyl-CoA dehydrogenase 7 NADH 21     
Isocitrate dehydrogenase 8 NADH 24     
α-Ketoglutarate dehydrogenase 8 NADH 24     
Succinyl-CoA synthetase 8*     
Succinate dehydrogenase 8 FADH2 16     
Malate dehydrogenase 8 NADH 24     
     Total 131     

* GTP produced directly in this step yields ATP in the reaction catalyzed by nucleoside
diphosphate kinase (p. 457).
Figure 16–10  The oxidation of a monounsaturated fatty acyl-CoA, such as oleoyl-CoA (Δ9), requires an additional enzyme, enoyl-CoA isomerase. This enzyme repositions the double bond, converting the cis isomer to a trans isomer, a normal intermediate in β oxidation.

The fatty acid oxidation sequence just described is typical when the incoming fatty acid is saturated (having only single bonds in its carbon chain). However, most of the fatty acids in the triacylglycerols and phospholipids of animals and plants are unsaturated, having one or more double bonds. These bonds are in the cis configuration and cannot be acted upon by enoyl-CoA hydratase, the enzyme catalyzing the addition of H2O to the trans double bond of the Δ2-enoyl-CoA generated during β oxidation. However, by the action of two auxiliary enzymes, the fatty acid oxidation sequence described above can also break down the common unsaturated fatty acids. The action of these two enzymes, one an isomerase and the other a reductase, will be illustrated by two examples.

First, let us follow the oxidation of oleate, an abundant 18-carbon monounsaturated fatty acid with a cis double bond between C-9 and C-10 (denoted Δ9). Oleate is converted into oleoyl-CoA (Fig. 16–10),

which is transported through the mitochondrial membrane as oleoyl-carnitine and then converted back into oleoyl-CoA in the matrix (Fig. 16–6). Oleoyl-CoA then undergoes three passes through the fatty acid oxidation cycle to yield three molecules of acetyl-CoA and the coenzyme A ester of a Δ3, 12-carbon unsaturated fatty acid, cis3-dodecenoyl-CoA (Fig. 16–10). This product cannot be acted upon by the next enzyme of the β-oxidation pathway, enoyl-CoA hydratase, which acts only on trans double bonds. However, by the action of the auxiliary enzyme, enoyl-CoA isomerase, the cis3-enoyl-CoA is isomerized to yield the trans2-enoyl-CoA, which is converted by enoyl-CoA hydratase into the corresponding L-β-hydroxyacyl-CoA (trans2-dodecenoyl-CoA). This intermediate is now acted upon by the remaining enzymes of β oxidation to yield acetyl-CoA and a 10-carbon saturated fatty acid as its coenzyme A ester (decanoyl-CoA). The latter undergoes four more passes through the pathway to yield altogether nine acetyl-CoAs from one molecule of the 18-carbon oleate.
Figure 16–11  Oxidation of polyunsaturated fatty acids requires a second auxiliary enzyme in addition to enoyl-CoA isomerase: NADPH-dependent 2,4-dienoyl-CoA reductase. The combined action of these two enzymes converts a trans2,cis4-dienoyl-CoA intermediate into the trans2-enoyl-CoA substrate necessary for β oxidation.

The other auxiliary enzyme (a reductase) is required for oxidation of polyunsaturated fatty acids. As an example, we take the 18-carbon linoleate, which has a cis9,cis12 configuration (Fig. 16–11). Linoleoyl-CoA undergoes three passes through the standard β-oxidation sequence to yield three molecules of acetyl-CoA and the coenzyme A ester of a 12-carbon unsaturated fatty acid with a cis3,cis6 configuration. This intermediate cannot be used by the enzymes of the β-oxidation pathway; its double bonds are in the wrong position and have the wrong configuration (cis, not trans). However, the combined action of enoyl-CoA isomerase and 2,4-dienoyl-CoA reductase (Fig. 16–11) allows reentry of this intermediate into the normal β-oxidation pathway and its degradation to six acetyl-CoAs. The overall result is conversion of linoleate to nine molecules of acetyl-CoA.

Although most naturally occurring lipids contain fatty acids with an even number of carbon atoms, fatty acids with an odd number of carbons are found in significant amounts in the lipids of many plants and some marine organisms. Small quantities of the three-carbon propionate are added as a mold inhibitor to some breads and cereals, and thus propionate enters the human diet. Moreover, cattle and other ruminant animals form large amounts of propionate during fermentation of carbohydrates in the rumen. The propionate so formed is absorbed into the blood and oxidized by the liver and other tissues.

Figure 16–12  Strategy for the oxidation of propionyl-CoA, involving the carboxylation of propionyl-CoA to D-methylmalonyl-CoA and conversion of the latter to succinyl-CoA. This conversion requires epimerization of D- to L-methylmalonyl-CoA, followed by a remarkable reaction in which substituents on adjacent carbon atoms exchange positions; Box 16–2 describes the role of coenzyme B12 in this exchange reaction.

Long-chain odd-carbon fatty acids are oxidized by the same pathway as the even-carbon acids, beginning at the carboxyl end of the chain. However, the substrate for the last pass through the β-oxidation sequence is a fatty acyl-CoA in which the fatty acid has five carbon

atoms. When this is oxidized and ultimately cleaved, the products are acetyl-CoA and propionyl-CoA. The acetyl-CoA is of course oxidized via the citric acid cycle, but propionyl-CoA takes a rather unusual enzymatic pathway, involving three enzymes. Propionyl-CoA is carboxylated to form the D stereoisomer of methylmalonyl-CoA (Fig. 16–12) by propionyl-CoA carboxylase, which contains the cofactor biotin. In this enzymatic reaction, as in the pyruvate carboxylase reaction (see Fig. 15–13), CO2 (or its hydrated ion, HCO3 ) is activated by attachment to biotin before its transfer to the propionate moiety. The formation of the carboxybiotin intermediate requires energy, which is provided by the cleavage of ATP to ADP and Pi.

The D-methylmalonyl-CoA thus formed is enzymatically epimerized to its L stereoisomer by the action of methylmalonyl-CoA epimerase (Fig. 16–12). The L-methylmalonyl-CoA undergoes an intramolecular rearrangement to form succinyl-CoA, which can enter the citric acid cycle. This rearrangement is catalyzed by methylmalonyl-CoA mutase, which requires as its coenzyme deoxyadenosylcobalamin, or coenzyme B12, derived from vitamin B12 (cobalamin) (Box 16–2).

B O X  16–2
Coenzyme B12: A Radical Solution to a Perplexing Problem

In the methylmalonyl-CoA mutase reaction (see Fig. 16–12), the group –CO–S-CoA at C-2 of the original propionate exchanges position with a hydrogen atom at C-3 of the original propionate (Fig. 1a). Coenzyme B12 is the cofactor for this reaction, as it is for almost all enzymes that catalyze reactions of this general type (Fig. 1b). These coenzyme B12-dependent reactions are among the very few enzymatic reactions in biology in which there is an exchange of an alkyl or substituted alkyl group (X) with a hydrogen atom of an adjacent carbon, with no mixing of the hydrogen atom transferred with the hydrogen of the solvent, H2O. How is it possible for the hydrogen atom to move between two carbons without mixing with the enormous excess of hydrogen atoms in the solvent?

Coenzyme B12 is the cofactor form of vitamin B12, which is unique among all the vitamins in that it contains not only a complex organic molecule but also an essential trace element, cobalt. The complex corrin ring system of vitamin B12 (colored blue in Fig. 2), to which cobalt (as Co3+) is coordinated, is chemically related to the porphyrin ring system of heme and heme proteins (see Fig. 7–18). A fifth coordination position of cobalt is filled by a nucleotide we have not encountered before, dimethylbenzimidazole ribonucleotide (yellow), bound covalently by its 3′-phosphate group to one of the side chains of the corrin ring through aminoisopropanol.

Vitamin B12 as usually isolated is called cyanocobalamin because it contains a cyano group (picked up during purification) attached to cobalt in the sixth coordination position. In 5′-deoxyadenosylcobalamin, the cofactor for methylmalonyl-CoA mutase, the cyano group is replaced by the 5′-deoxyadenosyl group (red in Fig. 2), covalently bound through C-5′ to the cobalt. The three-dimensional structure of the cofactor was determined by x-ray crystallography by Dorothy Crowfoot Hodgkin in 1956.

The formation of this complex cofactor (Fig. 3) is one of only two known cases in which triphosphate is cleaved from ATP; the other case is the formation of S-adenosylmethionine from ATP and methionine (see Fig. 17–20).

The key to understanding how coenzyme B12 catalyzes hydrogen exchange lies in the properties of the covalent bond between cobalt and C-5′ of the deoxyadenosyl group (Fig. 2). This is a relatively weak bond; its bond dissociation energy is about 110 kJ/mol, compared with 348 kJ/mol for a typical C–C bond or 414 kJ/mol for a C–H bond. Merely illuminating the compound with visible light is enough to break this bond.
(This extreme photolability probably accounts for the fact that plants do not contain vitamin B12.) Dissociation produces a 5′-deoxyadenosyl radical and the Co2+ form of the vitamin. The chemical function of 5′-deoxyadenosylcobalamin is to generate free radicals in this way, initiating a series of transformations such as that illustrated in Figure 4, a postulated mechanism for the reaction catalyzed by methylmalonyl-CoA mutase and a number of other coenzyme B12-dependent transformations.

The enzyme first breaks the Co–C bond in the cofactor, leaving the coenzyme in its Co2+ form and producing the 5′-deoxyadenosyl free radical (step ). This radical now abstracts a hydrogen atom from the substrate, converting the substrate to a radical and producing 5′-deoxyadenosine (step ). Rearrangement of the substrate radical (step ) yields another radical, in which the migrating

group X (–CO–S-CoA for methylmalonyl-CoA mutase) has moved to the adjacent carbon to form a productlike radical. The hydrogen atom initially abstracted from the substrate is now part of the CH3– group of 5′-deoxyadenosine; one of the hydrogens from this same CH3– group (it can be the same one originally abstracted) is returned to the productlike radical, generating the product and regenerating the deoxyadenosyl free radical (step ). Finally, the bond re-forms between cobalt and the CH2– group of the deoxyadenosyl radical (step ), destroying the free radical and regenerating the cofactor in its Co3+ form, ready to undergo another catalytic cycle.

In this postulated mechanism, the migrating hydrogen atom never exists as a free species and is thus never free to exchange with the hydrogen of surrounding water molecules.

Vitamin B12 deficiency results in serious disease. Vitamin B12 is not made by either plants or animals and can be synthesized by only a few species of microorganisms. It is required in only minute amounts, about 3 μg/day, by healthy people, but the severe disease pernicious anemia results from failure to absorb vitamin B12 efficiently from the intestine, where it is synthesized by intestinal bacteria or obtained from digestion of meat in the diet. A glycoprotein essential to vitamin B12 absorption, called intrinsic factor, is not produced in sufficient quantity in individuals with this disease. The pathology in pernicious anemia includes reduced production of erythrocytes, reduced levels of hemoglobin, and severe, progressive impairment of the central nervous system. Administration of large doses of vitamin B12 alleviates these symptoms in at least some cases.

In the liver, fatty acyl–CoAs formed in the cytosol have two major pathways open to them: (1) β oxidation by enzymes in the mitochondria or (2) conversion into triacylglycerols and phospholipids by enzymes in the cytosol. The pathway taken depends upon the rate of transfer of long-chain fatty acyl–CoAs into the mitochondria. The three-step process by which fatty acyl groups are carried from cytosolic fatty acyl–CoA into the mitochondrial matrix (Fig. 16–6) is rate-limiting for fatty acid oxidation. Once fatty acyl groups have entered the mitochondria, they are committed to oxidation to acetyl-CoA.

Malonyl-CoA, the first intermediate in the cytosolic biosynthesis of long-chain fatty acids from acetyl-CoA (Chapter 20), increases in concentration whenever the animal is well supplied with carbohydrate; excess glucose that cannot be oxidized or stored as glycogen is converted in the cytosol into fatty acids for storage as triacylglycerol. The inhibition of carnitine acyltransferase I by malonyl-CoA assures that the oxidation of fatty acids is inhibited whenever the liver is amply supplied with glucose as fuel and is actively making triacylglycerols from excess glucose.

Two of the enzymes of β oxidation are also regulated by metabolites that signal energy sufficiency. When the [NADH]/[NAD+] ratio is high, β-hydroxyacyl-CoA dehydrogenase is inhibited; in addition, high concentrations of acetyl-CoA inhibit thiolase.

Figure 16–13  Comparison of β oxidation of fatty acids as it occurs in animal mitochondria and in animal and plant peroxisomes. The peroxisomal system differs in two respects: (1) in the first oxidative step electrons pass directly to O2, generating H2O2, and (2) the NADH formed in β oxidation cannot be reoxidized, and the peroxisome must export reducing equivalents to the cytosol. (These eventually are passed on to mitochondria.) Fatty acid oxidation in glyoxysomes occurs by the peroxisomal pathway. In mitochondria, acetyl-CoA is further oxidized via the citric acid cycle. Acetyl-CoA produced by peroxisomes and glyoxysomes is exported; the acetate from glyoxysomes serves as a biosynthetic precursor (see Fig. 16–14).

Although the major site of fatty acid oxidation in animal cells is the mitochondrial matrix, other compartments in certain cells also contain enzymes capable of oxidizing fatty acids to acetyl-CoA, by a pathway similar to, but not identical with, that in mitochondria. Peroxisomes

are membrane-enclosed cellular compartments (see p. 38) in animals and plants, where hydrogen peroxide is produced by fatty acid oxidation and then destroyed enzymatically. As in the oxidation of fatty acids in mitochondria, the intermediates are coenzyme A derivatives, and the process consists of four steps (Fig. 16–13): (1) dehydrogenation; (2) addition of water to the resulting double bond; (3) oxidation of the β-hydroxyacyl-CoA to a ketone, and (4) thiolytic cleavage by coenzyme A. The difference between the peroxisomal and mitochondrial pathways is in the first step. In peroxisomes, the flavoprotein dehydrogenase that introduces the double bond passes electrons directly to O2, producing H2O2 (Fig. 16–13). This strong and potentially damaging oxidant is immediately cleaved to H2O and O2 by catalase. By contrast, in mitochondria the electrons removed in the first oxidation step pass through the respiratory chain to O2, and H2O is the product, a process accompanied by ATP synthesis. In peroxisomes, the energy released in the first oxidative step of fatty acid breakdown is dissipated as heat.

High concentrations of fats in the diet result in increased synthesis of the enzymes of peroxisomal β oxidation in mammalian liver. Liver peroxisomes do not contain the enzymes of the citric acid cycle and cannot catalyze the oxidation of acetyl-CoA to CO2. Instead, the acetate produced by fatty acid oxidation is exported from peroxisomes. Presumably some of this acetate enters mitochondria and is oxidized there.

Figure 16–14  The role of β oxidation in the conversion of seed triacylglycerols into glucose in germinating seeds.

Fatty acid oxidation in plants occurs in the peroxisomes of leaf tissue and the glyoxysomes of germinating seeds. Plant peroxisomes and glyoxysomes are similar in structure and function. Glyoxysomes occur only during seed germination, and may be considered specialized peroxisomes.

In plants, the biological role of β oxidation in peroxisomes and glyoxysomes is clear: it provides biosynthetic precursors from stored lipids. The β-oxidation pathway is not an important source of metabolic energy in plants; in fact, plant mitochondria do not contain the enzymes of β oxidation. During germination, triacylglycerols stored in seeds are converted into glucose and a wide variety of essential metabolites (Fig. 16–14). Fatty acids released from triacylglycerols are activated to their coenzyme A derivatives and oxidized in glyoxysomes by the same four-step process that occurs in peroxisomes (Fig. 16–13). The acetyl-CoA produced is converted via the glyoxylate cycle (Chapter 15) to four-carbon precursors for gluconeogenesis (Chapter 19). Glyoxysomes, like peroxisomes, contain high concentrations of catalase, which converts the H2O2 produced by β oxidation to H2O and O2.

Figure 16–15  A schematic diagram of the structure of the enzymes of β oxidation in gram-negative (a) and gram-positive (b) bacteria. The complex of four enzyme activities, two of which are part of a single polypeptide chain, is typical of gram-negative bacteria and is also found in the peroxisomal and glyoxysomal β-oxidation systems (a). The four enzymes of β oxidation in mitochondria are separate entities, similar to those of gram-positive bacteria (b). Enz1, acyl-CoA dehydrogenase; Enz2, enoyl-CoA hydratase; Enz3, L-β-hydroxyacyl-CoA dehydrogenase; Enz4, thiolase.

Although the β-oxidation reaction sequence in mitochondria is essentially the same as that in peroxisomes and glyoxysomes, the mitochondrial enzymes differ significantly from their isozymes in those compartments. These differences apparently reflect an evolutionary divergence that occurred very early, with the separation of gram-positive and gram-negative bacteria (see Fig. 2–6). In mitochondria, each of the four enzymes of β oxidation is a separate, soluble protein, similar in structure to the analogous enzyme in gram-positive bacteria. In contrast, the enzymes of peroxisomes and glyoxysomes are part of a complex of proteins, at least one of which contains two enzymatic activities in a single polypeptide chain (Fig. 16–15). Enoyl-CoA hydratase and

L-β-hydroxyacyl-CoA dehydrogenase activities both reside in a single, monomeric protein (Mr ~150,000), closely similar to the bifunctional protein of the gram-negative bacterium E. coli. The evolutionary selective value of retaining both types of β-oxidation system in the same organism is not yet apparent.

In human beings and most other mammals, acetyl-CoA formed in the liver during oxidation of fatty acids may enter the citric acid cycle (stage 2 of Fig. 16–7) or it may be converted to the “ketone bodies” acetoacetate, D-β-hydroxybutyrate, and acetone for export to other tissues. (The term “bodies” is a historical artifact; these compounds are soluble in blood and urine.) Acetone, produced in smaller quantities than the other ketone bodies, is exhaled. Acetoacetate and D-β-hydroxybutyrate are transported by the blood to the extrahepatic tissues, where they are oxidized via the citric acid cycle to provide much of the energy required by tissues such as skeletal and heart muscle and the renal cortex. The brain, which normally prefers glucose as a fuel, can adapt to the use of acetoacetate or D-β-hydroxybutyrate under starvation conditions, when glucose is unavailable.

A major determinant of the pathway taken by acetyl-CoA in liver mitochondria is the availability of oxaloacetate to initiate entry of acetyl-CoA into the citric acid cycle. Under some circumstances (such as starvation) oxaloacetate is drawn out of the citric acid cycle for use in synthesizing glucose. When the oxaloacetate concentration is very low, little acetyl-CoA enters the cycle, and ketone body formation is favored. The production and export of ketone bodies from the liver to extrahepatic tissues allows continued oxidation of fatty acids in the liver when acetyl-CoA is not being oxidized via the citric acid cycle. Overproduction of ketone bodies can occur in conditions of severe starvation and in uncontrolled diabetes.

Figure 16–16  Formation of ketone bodies from acetyl-CoA. Under circumstances that cause acetyl-CoA accumulation (starvation or untreated diabetes, for example), thiolase catalyzes the condensation of two acetyl-CoA molecules to acetoacetyl-CoA, the parent of the three ketone bodies. These reactions all occur within the mitochondrial matrix. The six-carbon compound β-hydroxy-βmethylglutaryl-CoA (HMG-CoA) is also an intermediate of sterol biosynthesis, but the enzyme that forms HMG-CoA in that pathway is cytosolic. HMG-CoA lyase is present in the mitochondrial matrix but not in the cytosol.

The first step in formation of acetoacetate in the liver (Fig. 16–16) is the enzymatic condensation of two molecules of acetyl-CoA, catalyzed by thiolase; this is simply the reversal of the last step of β oxidation. The acetoacetyl-CoA then condenses with acetyl-CoA to form β-hydroxy-β-methylglutaryl-CoA (HMG-CoA), which is cleaved to free acetoacetate and acetyl-CoA.

The free acetoacetate so produced is reversibly reduced by D-β-hydroxybutyrate dehydrogenase, a mitochondrial enzyme, to D-β-hydroxybutyrate (Fig. 16–16). This enzyme is specific for the D stereoisomer; it does not act on L-β-hydroxyacyl-CoAs and is not to be confused with L-β-hydroxyacyl-CoA dehydrogenase, which acts in the β-oxidation pathway. In healthy people, acetone is formed in very small amounts from acetoacetate by the loss of a carboxyl group. Acetoacetate is easily decarboxylated; the carboxyl group may be lost spontaneously or by the action of acetoacetate decarboxylase (Fig. 16–16). Because untreated diabetics produce large quantities of acetoacetate, their blood contains significant amounts of acetone, which is toxic. Acetone is volatile and imparts a characteristic odor to the breath, which is sometimes useful in diagnosing the severity of the disease.

Figure 16–17  Hydroxybutyrate as a fuel. D-β-Hydroxybutyrate synthesized in the liver passes into the blood and thus to other tissues, where it is converted to acetyl-CoA for energy production. It is first oxidized to acetoacetate, which is activated with coenzyme A donated from succinyl-CoA, then split by thiolase.

In the extrahepatic tissues D-β-hydroxybutyrate is oxidized to acetoacetate by D-β-hydroxybutyrate dehydrogenase (Fig. 16–17). Acetoacetate is activated to form its coenzyme A ester by transfer of CoA from succinyl-CoA, an intermediate of the citric acid cycle (see Fig. 15–7), in a reaction catalyzed by β-ketoacyl-CoA transferase. The acetoacetyl-CoA is then cleaved by thiolase to yield two acetyl-CoAs, which enter the citric acid cycle.

Figure 16–18  Ketone body formation and export from the liver. Conditions that increase gluconeogenesis (diabetes, fasting) slow the citric acid cycle (by drawing off oxaloacetate) and enhance the conversion of acetyl-CoA to acetoacetate. The released coenzyme A allows continued β oxidation of fatty acids.

The production and export of ketone bodies from the liver allows continued oxidation of fatty acids with only minimal oxidation of acetyl-CoA in the liver (Fig. 16–18). When, for example, intermediates of the citric acid cycle are being used for glucose synthesis via gluconeogenesis, oxidation of citric acid cycle intermediates slows, and so does acetyl-CoA oxidation. Moreover, the liver contains a limited amount of coenzyme A, and when most of it is tied up in acetyl-CoA, β oxidation of fatty acids slows for lack of the free coenzyme. The production and export of ketone bodies frees coenzyme A, allowing continued fatty acid oxidation.

Severe starvation or untreated diabetes mellitus leads to overproduction of ketone bodies, with several associated medical problems. During starvation, gluconeogenesis depletes citric acid cycle intermediates, diverting acetyl-CoA to ketone body production (Fig. 16–18). In untreated diabetes, insulin is present in insufficient quantity, and the extrahepatic tissues cannot take up glucose efficiently from the blood (Chapter 22). To raise the blood glucose level, gluconeogenesis in the liver accelerates, as does fatty acid oxidation in liver and muscle, with the result that ketone bodies are produced beyond the capacity of extrahepatic tissues to oxidize them. The rise in blood levels of acetoacetate and D-β-hydroxybutyrate lowers the blood pH, causing the condition known as acidosis. Extreme acidosis can lead to coma and in some cases death. Ketone bodies in the blood and urine of untreated diabetics may reach extraordinary levels (Table 16–2); this condition is ketosis. In individuals on very low-calorie diets, fats stored in adipose tissue become the major energy source. The levels of ketone bodies in the blood and urine should be monitored to avoid the dangers of acidosis and ketosis (ketoacidosis).
Figure 17–1  Overview of the catabolism of amino acids. The separate paths taken by the carbon skeleton and the amino groups are emphasized by the orange divergent arrow.
Chapter 17
Amino Acid Oxidation and the Production of Urea

Amino acids, derived largely from protein in the diet or from degradation of intracellular proteins, are the final class of biomolecules whose oxidation makes a significant contribution to the generation of metabolic energy. The fraction of metabolic energy derived from amino acids varies greatly with the type of organism and with the metabolic situation in which an organism finds itself. Carnivores, immediately following a meal, may obtain up to 90% of their energy requirements from amino acid oxidation. Herbivores may obtain only a small fraction of their energy needs from this source. Most microorganisms can scavenge amino acids from their environment if they are available; these can be oxidized as fuel when required by metabolic conditions. Photosynthetic plants, on the other hand, rarely, if ever, oxidize amino acids to provide energy. Instead, they convert CO2 and H2O into the carbohydrate that is used almost exclusively as an energy source. The amounts of amino acids in plant tissues are carefully regulated to just meet the requirements for biosynthesis of proteins, nucleic acids, and a few other molecules needed to support growth. Amino acid catabolism does occur in plants, but it is generally concerned with the production of metabolites for other biosynthetic pathways.

In animals, amino acids can undergo oxidative degradation in three different metabolic circumstances. (1) During the normal synthesis and degradation of cellular proteins (protein turnover; Chapter 26) some of the amino acids released during protein breakdown will undergo oxidative degradation if they are not needed for new protein synthesis. (2) When a diet is rich in protein, and amino acids are ingested in excess of the body's needs for protein synthesis, the surplus may be catabolized; amino acids cannot be stored. (3) During starvation or in diabetes mellitus, when carbohydrates are either unavailable or not properly utilized, body proteins are called upon as fuel. Under these different circumstances, amino acids lose their amino groups, and the α-keto acids so formed may undergo oxidation to CO2 and H2O. In addition, and often equally important, the carbon skeletons of the amino acids provide three- and four-carbon units that can be converted to glucose, which in turn can fuel the functions of the brain, muscle, and other tissues.

Amino acid degradative pathways are quite similar in most organisms. The focus of this chapter is on vertebrates, because amino acid catabolism has received the most attention in these organisms. As is the case for sugar and fatty acid catabolic pathways, the processes of amino acid degradation converge on the central catabolic pathways for

carbon metabolism. The carbon skeletons of the amino acids generally find their way to the citric acid cycle, and from there they are either oxidized to produce chemical energy or funneled into gluconeogenesis. In some cases the reaction pathways closely parallel steps in the catabolism of fatty acids (Chapter 16).

However, one major factor distinguishes amino acid degradation from the catabolic processes described to this point: every amino acid contains an amino group. Every degradative pathway therefore passes through a key step in which the α-amino group is separated from the carbon skeleton and shunted into the specialized pathways for amino group metabolism (Fig. 17–1). This biochemical fork in the road is the point around which this chapter is organized. We deal first with amino group metabolism and nitrogen excretion, then with the fate of the carbon skeletons derived from the amino acids.

Figure 17–2  Overview of amino group catabolism in the vertebrate liver (shaded).
Excess NH4+ is excreted as urea or uric acid.

Nitrogen ranks fourth, behind carbon, hydrogen, and oxygen, in its contribution to the mass of living cells. Atmospheric nitrogen, N2, is abundant but is too inert for use in most biochemical processes. Because only a few microorganisms can convert N2 to biologically useful forms such as NH3, amino groups are used with great economy in biological systems.

An overview of the catabolism of ammonia and amino groups in vertebrates is provided in Figure 17–2 (p. 508). Amino acids derived from dietary proteins are the source of most amino groups. Most of the amino acids are metabolized in the liver. Some of the ammonia that is generated is recycled and used in a variety of biosynthetic processes; the excess is either excreted directly or converted to uric acid or urea for excretion, depending on the organism. Excess ammonia generated in other (extrahepatic) tissues is transported to the liver (in the form of amino groups, as described below) for conversion to the appropriate excreted form. With these reactions we encounter the coenzyme pyridoxal phosphate, the functional form of vitamin B6 and a coenzyme of major importance in nitrogen metabolism.

The amino acids glutamate and glutamine play especially critical roles in these pathways (Fig. 17–2). Amino groups from amino acids are generally first transferred to α-ketoglutarate in the cytosol of liver cells (hepatocytes) to form glutamate. Glutamate is then transported into the mitochondria; only here is the amino group removed to form NH4+. Excess ammonia generated in most other tissues is converted to the amide nitrogen of glutamine, then transported to liver mitochondria. In most tissues, one or both of these amino acids are found in elevated concentrations relative to other amino acids.

In muscle, excess amino groups are generally transferred to pyruvate to form alanine. Alanine is another important molecule in the transport of amino groups, conveying them from muscle to the liver.

We begin with a discussion of the breakdown of dietary proteins to amino acids, then turn to a general description of the metabolic fates of amino groups.

Figure 17–3  A portion of the human digestive tract. (a) Gastric glands in the stomach lining. The parietal cells and chief cells secrete their products in response to the hormone gastrin.
(b) Exocrine cells of the pancreas. The cytoplasm is completely filled with rough endoplasmic reticulum, on which the ribosomes synthesize the polypeptide chains of the zymogens of many digestive enzymes. The zymogens are concentrated in condensing vesicles, ultimately forming mature zymogen granules. When the cell is stimulated, the plasma membrane fuses with the membrane around the zymogen granules, and the zymogens are released into the lumen of the collecting duct by exocytosis. The collecting ducts ultimately lead to the pancreatic duct and thence to the small intestine. (c) Villi of the small intestine. Amino acids are absorbed through the epithelial cell layer (intestinal mucosa) and enter the capillaries. Recall that the products of lipid hydrolysis in the small intestine enter the lymphatic system following absorption by the intestinal mucosa (Fig. 16–1).

In humans, the degradation of ingested proteins into their constituent amino acids occurs in the gastrointestinal tract. Entry of protein into the stomach stimulates the gastric mucosa to secrete the hormone gastrin, which in turn stimulates the secretion of hydrochloric acid by the parietal cells of the gastric glands (Fig. 17–3a) and pepsinogen by the chief cells. The acidity of gastric juice (pH 1.5 to 2.5) acts as an antiseptic and kills most bacteria and other foreign cells. Globular proteins denature at low pH, rendering their internal peptide bonds more accessible to enzymatic hydrolysis. Pepsinogen (Mr 40,000), an inactive precursor or zymogen (p. 235), is converted into active pepsin in the gastric juice by the enzymatic action of pepsin itself. In this process, 42 amino acid residues are removed from the amino-terminal end of the polypeptide chain. The portion of the molecule that remains intact is enzymatically active pepsin (Mr 33,000). In the stomach, pepsin hydrolyzes ingested proteins at peptide bonds on the amino-terminal side of the aromatic amino acid residues Tyr, Phe, and Trp (see Table 6–7), cleaving long polypeptide chains into a mixture of smaller peptides.

As the acidic stomach contents pass into the small intestine, the low pH triggers the secretion of the hormone secretin into the blood. Secretin stimulates the pancreas to secrete bicarbonate into the small intestine to neutralize the gastric HCl, increasing the pH abruptly to about pH 7. The digestion of proteins continues in the small intestine. The entry of amino acids into the upper part of the intestine (duodenum) releases the hormone cholecystokinin, which stimulates secretion of several pancreatic enzymes, whose activity optima occur at pH 7 to 8. Three of these, trypsin, chymotrypsin, and carboxypeptidase, are made by the exocrine cells of the pancreas (Fig. 17–3b) as their respective enzymatically inactive zymogens, trypsinogen, chymotrypsinogen, and procarboxypeptidase.

Synthesis of these enzymes as inactive precursors protects the exocrine cells from destructive proteolytic attack. The pancreas protects itself against self digestion in another way – by making a specific inhibitor, itself a protein, called pancreatic trypsin inhibitor (p. 235). Free trypsin can activate not only trypsinogen but also three other digestive zymogens: chymotrypsinogen, procarboxypeptidase, and proelastase; trypsin inhibitor effectively prevents premature production of free proteolytic enzymes within the pancreatic cells.

After trypsinogen enters the small intestine, it is converted into its active form, trypsin, by enteropeptidase, a specialized proteolytic enzyme secreted by intestinal cells. Once some free trypsin has been formed, it also can catalyze the conversion of trypsinogen into trypsin (see Fig. 8–30). Trypsin, as noted above, can convert chymotrypsinogen and procarboxypeptidase into chymotrypsin and carboxypeptidase.

Trypsin and chymotrypsin thus hydrolyze into smaller peptides the peptides resulting from the action of pepsin in the stomach. This stage of protein digestion is accomplished very efficiently because pepsin, trypsin, and chymotrypsin have different amino acid specificities. Trypsin hydrolyzes those peptide bonds whose carbonyl groups are contributed by Lys and Arg residues, and chymotrypsin hydrolyzes peptide bonds on the carboxyl-terminal side of Phe, Tyr, and Trp residues (see Table 6–7).

Degradation of the short peptides in the small intestine is now completed by other peptidases. The first is carboxypeptidase, a zinc-containing enzyme, which removes successive carboxyl-terminal residues from peptides. The small intestine also secretes an aminopeptidase, which can hydrolyze successive amino-terminal residues from short peptides. By the sequential action of these proteolytic enzymes and peptidases, ingested proteins are hydrolyzed to yield a mixture of free amino acids, which can then be transported across the epithelial cells lining the small intestine (Fig. 17–3c). The free amino acids enter the blood capillaries in the villi and are transported to the liver.

In humans, most globular proteins from animal sources are almost completely hydrolyzed into amino acids, but some fibrous proteins, such as keratin, are only partially digested. Many proteins of plant foods, such as cereal grains, are incompletely digested because the protein part of grains of seeds is surrounded by indigestible cellulose husks.

Celiac disease is a condition in which the intestinal enzymes are unable to digest certain water-insoluble proteins of wheat, particularly gliadin, which is injurious to the cells lining the small intestine. Wheat products must therefore be avoided by such individuals. Another disease involving the proteolytic enzymes of the digestive tract is acute pancreatitis. In this condition, caused by obstruction of the normal

pathway of secretion of pancreatic juice into the intestine, the zymogens of the proteolytic enzymes are converted into their catalytically active forms prematurely, inside the pancreatic cells. As a result these powerful enzymes attack the pancreatic tissue itself, causing a painful and serious destruction of the organ, which can be fatal.
Figure 17–4  Excretory forms of amino group nitrogen in different forms of life. Notice that the carbon atoms of urea and uric acid are at a high oxidation state; the organism discards carbon only after having obtained most of its available energy of oxidation.

The α-amino groups of the 20 L-amino acids commonly found in proteins are removed during the oxidative degradation of the amino acids. If not reused for synthesis of new amino acids or other nitrogenous products, these amino groups are channeled into a single excretory end product (Fig. 17–4). Many aquatic organisms simply release ammonia as NH4+ into the surrounding medium. Most terrestrial vertebrates first convert the ammonia into urea (humans, other mammals, and adult amphibians) or uric acid (birds, reptiles).

Figure 17–5  (a) The aminotransferase reaction (transamination). In many aminotransferase reactions, α-ketoglutarate is the amino group acceptor. All aminotransferases have pyridoxal phosphate (PLP) as cofactor. (b) The reaction of alanine aminotransferase is shown as an example.
The removal of the α-amino groups, the first step in the catabolism of most of the L-amino acids, is promoted by enzymes called aminotransferases or transaminases. In these transamination reactions, the α-amino group is transferred to the α-carbon atom of α-ketoglutarate, leaving behind the corresponding α-keto acid analog of the amino acid (Fig. 17–5). There is no net deamination (i.e., loss of amino groups) in such reactions because the α-ketoglutarate becomes aminated as the α-amino acid is deaminated. The effect of transamination reactions is to collect the amino groups from many different amino acids in the form of only one, namely, L-glutamate. The glutamate channels amino groups either into biosynthetic pathways or into a final sequence of reactions by which nitrogenous waste products are formed and then excreted.

Cells contain several different aminotransferases, many specific for α-ketoglutarate as the amino group acceptor. The aminotransferases differ in their specificity for the other substrate, the L-amino acid that donates the amino group, and are named for the amino group donor (Fig. 17–5b). The reactions catalyzed by the aminotransferases are freely reversible, having an equilibrium constant of about 1.0 (ΔG°’ ≈ 0 kJ/mol).

Figure 17–6  The prosthetic group of aminotransferases. (a) Pyridoxal phosphate (PLP) and its aminated form pyridoxamine phosphate are the tightly bound coenzymes of aminotransferases. The functional groups involved in their action are shaded in red. Pyridoxal phosphate is bound to the enzyme both through strong noncovalent interactions and through formation of a Schiff base linkage involving a Lys residue at the active site (b).
All aminotransferases share a common prosthetic group and a common reaction mechanism. The prosthetic group is pyridoxal phosphate (PLP), the coenzyme form of pyridoxine or vitamin B6. Pyridoxal phosphate was briefly introduced in Chapter 14 (p. 422) as a cofactor in the glycogen phosphorylase reaction. Its role in that reaction, however, is not representative of its normal coenzyme function. Its more typical functions occur in the metabolism of molecules with amino groups.

Pyridoxal phosphate functions as an intermediate carrier of amino groups at the active site of arninotransferases. It undergoes reversible transformations between its aldehyde form, pyridoxal phosphate, which can accept an amino group, and its aminated form, pyridoxamine phosphate, which can donate its amino group to an α-keto acid (Fig. 17–6a). Pyridoxal phosphate is generally bound covalently to the enzyme’s active site through an imine (Schiff base) linkage to the ϵ-amino group of a Lys residue (Fig. 17–6b).

Pyridoxal phosphate is involved in a variety of reactions at the α and β carbons of amino acids. Reactions at the α carbon (Fig. 17–7) include racemizations (interconverting L- and D-amino acids) and decarboxylations, as well as transaminations. Pyridoxal phosphate plays the same chemical role in each of these reactions. One of the bonds to the a carbon is broken, removing either a proton or a carboxyl group and leaving behind a free electron pair on the carbon (a carbanion). This intermediate is very unstable and normally would not form at a significant rate. Pyridoxal phosphate provides a highly conjugated structure (an electron sink) that permits delocalization of the negative charge, stabilizing the carbanion (Fig. 17–7).

Aminotransferases are classic examples of enzymes catalyzing bimolecular ping-pong reactions (see Fig. 8–13b). In such reactions the first substrate must leave the active site before the second substrate can bind. Thus the incoming amino acid binds to the active site,

donates its amino group to pyridoxal phosphate, and departs in the form of an α-keto acid. Then the incoming α-keto acid is bound, accepts the amino group from pyridoxamine phosphate, and departs in the form of an amino acid.

The measurement of alanine aminotransferase and aspartate aminotransferase levels in blood serum is an important diagnostic procedure in medicine, used as an indicator of heart damage and to monitor recovery from the damage (Box 17–1).

Figure 17–7  Some of the amino acid transformations facilitated by pyridoxal phosphate. Pyridoxal phosphate is generally bound to the enzyme by means of a Schiff base (see Fig. 17–6b). Reactions begin with formation of a new Schiff base (aldimine) between the α-amino group of the amino acid and PLP, which substitutes for the enzyme–PLP linkage. The amino acid then can have three alternative fates, each involving formation of a carbanion: transamination, racemization, or decarboxylation. The Schiff base formed between PLP and the amino acid is in conjugation with the pyridine ring, which acts as an electron sink, permitting delocalization of the negative charge of the carbanion (as shown within the brackets). A quinonoid intermediate is involved in all of the reactions. The transamination route is especially important in the pathways described in this chapter. The highlighted transamination pathway (shown left to right) represents only part of the reaction catalyzed by aminotransferases. To complete the process, a second α-keto acid replaces the one that is released and is converted to an amino acid in a reversal of the reaction (right to left).

B O X  17–1
Assays for Tissue Damage

Analysis of different enzyme activities in blood serum gives valuable diagnostic information for a number of disease conditions.

Alanine aminotransferase (ALT; also called glutamate–pyruvate transaminase, GPT) and aspartate aminotransferase (AST; also called glutamate–oxaloacetate transaminase, GOT) are important in the diagnosis of heart and liver damage. Occlusion of a coronary artery by lipid deposits can cause severe local oxygen starvation and ultimately the degeneration of a localized portion of the heart muscle; this process is called myocardial infarction. Such damage causes aminotransferases, among other enzymes, to leak from the injured heart cells into the bloodstream. Measurements of the concentration in the blood serum of these two aminotransferases by the SGPT and SGOT tests (S for serum) and of another heart enzyme, creatine kinase (the SCK test), can provide information about the severity and the stage of the damage to the heart. Creatine kinase is the first heart enzyme to appear in the blood after a heart attack; it also disappears quickly from the blood. GOT is the next to appear, and GPT follows later. Lactate dehydrogenase also leaks from injured or anaerobic heart muscle.

SGOT and SGPT are also important in industrial medicine to determine whether people exposed to carbon tetrachloride, chloroform, or other solvents used in the chemical, dry-cleaning, and other industries have suffered liver damage. These solvents cause liver degeneration, with resulting leakage into the blood of various enzymes from the injured hepatocytes. Aminotransferases, because they are very active in liver and their activity can be detected in very small amounts, are most useful in the monitoring of people exposed to such industrial chemicals.

Figure 17–8  The reaction catalyzed by glutamate dehydrogenase. This enzyme can employ either NAD+ or NADP+ as cofactor, and is allosterically regulated by GTP and ADP.

We have seen that, in the liver, amino groups are removed from many of the α-amino acids by transamination with α-ketoglutarate to form L-glutamate. How are amino groups removed from glutamate to prepare them for excretion?

Glutamate is transported from the cytosol to the mitochondria, where it undergoes oxidative deamination catalyzed by L-glutamate dehydrogenase (Mr 330,000). This enzyme, which is present only in the mitochondrial matrix, requires NAD+ (or NADP+) as the acceptor of the reducing equivalents (Fig. 17–8). The combined action of the aminotransferases and glutamate dehydrogenase is referred to as transdeamination. A few amino acids bypass the transdeamination pathway and undergo direct oxidative deamination. The fate of the NH4+ produced by either of these processes is discussed in detail later.

As might be expected from its central role in amino group metabolism, glutamate dehydrogenase is a complex allosteric enzyme. The

enzyme molecule consists of six identical subunits. It is influenced by the positive modulator ADP and by the negative modulator GTP, a product of the succinyl-CoA synthetase reaction in the citric acid cycle (p. 456). Whenever a hepatocyte needs fuel for the citric acid cycle, glutamate dehydrogenase activity increases, making α-ketoglutarate available for the citric acid cycle and releasing NH4+ for excretion. On the other hand, whenever GTP accumulates in the mitochondria as a result of high citric acid cycle activity, oxidative deamination of glutamate is inhibited.

Ammonia is quite toxic to animal tissues (we examine some possible reasons for this toxicity later). In most animals excess ammonia is converted into a nontoxic compound before export from extrahepatic tissues into the blood and thence to the liver or kidneys. Glutamate, which is so critical to intracellular amino group metabolism, is supplanted by L-glutamine for this transport function. In many tissues, including the brain, ammonia is enzymatically combined with glutamate to yield glutamine by the action of glutamine synthetase. This reaction requires ATP and occurs in two steps. In the first step, glutamate and ATP react to form ADP and a γ-glutamyl phosphate intermediate, which reacts with ammonia to produce glutamine and inorganic phosphate. We will encounter glutamine synthetase again in Chapter 21 when we consider nitrogen metabolism in microorganisms, where this enzyme serves as a portal for the entry of fixed nitrogen into biological systems. Glutamine is a nontoxic, neutral compound that can readily pass through cell membranes, whereas glutamate, which bears a net negative charge, cannot. In most land animals glutamine is carried in the blood to the liver. As is the case for the amino group of glutamate, the amide nitrogen is released as ammonia only within liver mitochondria, where the enzyme glutaminase converts glutamine to glutamate and NH4+.

Glutamine is a major transport form of ammonia; it is normally present in blood in much higher concentrations than other amino acids. In addition to its role in the transport of amino groups, glutamine serves as a source of amino groups in a variety of biosynthetic reactions (Chapter 21).

Figure 17–9  The glucose–alanine cycle. Alanine serves as a carrier of ammonia equivalents and of the carbon skeleton of pyruvate from muscle to liver. The ammonia is excreted, and the pyruvate is used to produce glucose, which is returned to the muscle.

Alanine also plays a special role in transporting amino groups to the liver in a nontoxic form, by the glucose–alanine cycle (Fig. 17–9). In muscle and certain other tissues that degrade amino acids for fuel, amino groups are collected in glutamate by transamination (Fig. 17–2). Glutamate may then be converted to glutamine for transport to the liver, or it may transfer its α-amino group to pyruvate, a readily available product of muscle glycolysis, by the action of alanine aminotransferase (Fig. 17–9). The alanine, with no net charge at pH near 7, passes into the blood and is carried to the liver. As with glutamine, excess nitrogen carried to the liver as alanine is eventually delivered as ammonia in the mitochondria. In a reversal of the alanine aminotransferase reaction described above, alanine transfers its amino group to α-ketoglutarate, forming glutamate in the cytosol. Some of this glutamate is transported into the mitochondria and acted on by glutamate dehydrogenase, releasing NH4+ (Fig. 17–8). Alternatively, transamination with oxaloacetate moves amino groups from glutamate to aspartate, another nitrogen donor in urea synthesis.

The use of alanine to transport ammonia from hard-working skeletal muscles to the liver is another example of the intrinsic economy of living organisms. Vigorously contracting skeletal muscles operate anaerobically, producing not only ammonia from protein breakdown but also large amounts of pyruvate from glycolysis. Both these products must find their way to the liver – ammonia to be converted into urea for excretion and pyruvate to be rebuilt into glucose and returned to the muscles. Animals thus solve two problems with one cycle: they move the carbon atoms of pyruvate, as well as excess ammonia, from muscle to liver as alanine. In the liver, alanine yields pyruvate, the starting material for gluconeogenesis, and releases NH4+ for urea synthesis. The energetic burden of gluconeogenesis is thus imposed on the liver rather than the muscle, so that the available ATP in the muscle can be devoted to muscle contraction.

The catabolic production of ammonia poses a serious biochemical problem because ammonia is very toxic. The molecular basis for this toxicity is not entirely understood. The terminal stages of ammonia intoxication in humans are characterized by the onset of a comatose state and other effects on the brain, so that much of the research and speculation has focused on this tissue. The major toxic effects of ammonia in brain probably involve changes in cellular pH and the depletion of certain citric acid cycle intermediates.

The protonated form of ammonia (ammonium ion) is a weak acid, and the unprotonated form is a strong base:

Most of the ammonia generated in catabolism is present as NH4+ at neutral pH. Although many of the reactions that produce ammonia, such as the glutamate dehydrogenase reaction, yield NH4+, a few reactions, such as that of adenosine deaminase (Chapter 21), produce NH3. Excessive amounts of NH3 cause alkalization of cellular fluids, which has complex effects on cellular metabolism.

Ridding the cytosol of excess ammonia involves reductive amination of α-ketoglutarate to form glutamate by glutamate dehydrogenase (the reverse of the reaction described earlier) and conversion of glutamate

to glutamine by glutamine synthetase. Both of these enzymes occur in high levels in the brain, although the second probably represents the more important pathway for removal of ammonia. The first reaction depletes cellular NADH and α-ketoglutarate required for ATP production in the cell. The second reaction depletes ATP itself. Overall, NH3 may interfere with the very high levels of ATP production required to maintain brain function.

Depletion of glutamate in the glutamine synthetase reaction may have additional effects on the brain. Glutamate, and the compound γ-aminobutyrate (GABA) that is derived from it (Chapter 21), are both important neurotransmitters; the sensitivity of the brain to ammonia may well reflect a depletion of neurotransmitters as well as changes in cellular pH and ATP metabolism.

As we close this discussion of amino group metabolism, note that we have described several processes that deposit excess ammonia in the mitochondria of hepatocytes (Fig. 17–2). We now turn to a discussion of the fate of that ammonia.

As already noted (Fig. 17–4), most aquatic species, such as the bony fishes, excrete amino nitrogen as ammonia and are thus called ammonotelic animals; most terrestrial animals excrete amino nitrogen in the form of urea and are thus ureotelic; and birds and reptiles excrete amino nitrogen as uric acid and are called uricotelic. Plants recycle virtually all amino groups, and nitrogen excretion occurs only under very unusual circumstances. There is no general pathway for nitrogen excretion in plants.

In ureotelic organisms, the ammonia in the mitochondria of hepatocytes is converted to urea via the urea cycle. This pathway was discovered in 1932 by Hans Krebs (who later also discovered the citric acid cycle) and a medical student associate, Kurt Henseleit. Urea production occurs almost exclusively in the liver, and it represents the fate of most of the ammonia that is channeled there. This pathway now becomes the focus of our discussion.

Figure 17–10  The urea cycle. The three amino acids found by Krebs and Henseleit to stimulate urea formation from ammonia in liver slices are boxed. As shown, ornithine and citrulline can serve as successive precursors of arginine. Note that citrulline and ornithine are nonstandard amino acids that are not found in proteins.

Using thin slices of liver suspended in a buffered aerobic medium, Krebs and Henseleit found that the rate of urea formation from ammonia was greatly accelerated by adding any one of three α-amino acids: ornithine, citrulline, or arginine. Each of these three compounds stimulated urea synthesis to a far greater extent than any of the other common nitrogenous compounds tested, and their structures suggested that they might be related in a sequence.

From these and other facts Krebs and Henseleit deduced that a cyclic process occurs (Fig. 17–10), in which ornithine plays a role resembling that of oxaloacetate in the citric acid cycle. A molecule of ornithine combines with one molecule of ammonia and one of CO2 to form citrulline. A second amino group is added to citrulline to form arginine, which is then hydrolyzed to yield urea, with regeneration of ornithine. Ureotelic animals have large amounts of the enzyme arginase in the liver. This enzyme catalyzes the irreversible hydrolysis of arginine to urea and ornithine. The ornithine is then ready for the next turn of the urea cycle. The urea is passed via the bloodstream to the kidneys and is excreted into the urine.

Figure 17–11  The urea cycle and the reactions that feed amino groups into it. Note that the enzymes catalyzing these reactions (named in the text) are distributed between the mitochondrial matrix and the cytosol. One amino group enters the urea cycle from carbamoyl phosphate (step ), formed in the matrix; the other (entering at step ) is derived from aspartate, also formed in the matrix via transamination of oxaloacetate and glutamate in a reaction catalyzed by aspartate aminotransferase. The urea cycle itself consists of four steps: Formation of citrulline from ornithine and carbamoyl phosphate. Citrulline passes into the cytosol. Formation of argininosuccinate through a citrullyl-AMP intermediate. Formation of arginine from argininosuccinate. This reaction releases fumarate, which enters the citric acid cycle. Formation of urea. The arginase reaction also regenerates the starting compound in the cycle, ornithine. The pathways by which NH4+ arrives in the mitochondrial matrix are discussed earlier in the text.

The urea cycle begins inside the mitochondria of hepatocytes, but three of the steps occur in the cytosol; the cycle thus spans two cellular compartments (Fig. 17–11). The first amino group to enter the urea cycle is derived from ammonia inside the mitochondria, arising by the multiple pathways described above. Some ammonia also arrives at the liver via the portal vein from the intestine, where it is produced by bacterial oxidation of amino acids. Whatever its source, the NH4+ generated in liver mitochondria is immediately used, together with HCO3 produced by mitochondrial respiration, to form carbamoyl phosphate in the matrix (Fig. 17–12; see also Fig. 17–11). This ATP-dependent reaction is catalyzed by the enzyme carbamoyl phosphate synthetase I. The mitochondrial form of the enzyme is distinct from the cytosolic (II) form, which has a separate function in pyrimidine biosynthesis (Chapter 21). Carbamoyl phosphate synthetase I is a regulatory enzyme; it requires N-acetylglutamate as a positive modulator (see below). Carbamoyl phosphate may be regarded as an activated carbamoyl group donor.

The carbamoyl phosphate now enters the urea cycle, which entails four enzymatic steps. Carbamoyl phosphate donates its carbamoyl group to ornithine to form citrulline and release Pi (Fig. 17–11, step )

in a reaction catalyzed by ornithine transcarbamoylase. The citrulline is released from the mitochondrion into the cytosol.

The second amino group is introduced from aspartate (generated in the mitochondria by transamination (Fig. 17–11) and transported to the cytosol) by a condensation reaction between the amino group of aspartate and the ureido (carbonyl) group of citrulline to form argininosuccinate (step ). This reaction, catalyzed by argininosuccinate synthetase of the cytosol, requires ATP and proceeds through a citrullyl-AMP intermediate. The argininosuccinate is then reversibly cleaved by argininosuccinate lyase to form free arginine and fumarate (step ), which enters the pool of citric acid cycle intermediates. In the last reaction of the urea cycle the cytosolic enzyme arginase cleaves arginine to yield urea and ornithine (step ). Ornithine is thus regenerated and can be transported into the mitochondrion to initiate another round of the urea cycle.

Figure 17–12  The reaction catalyzed by carbamoyl phosphate synthetase I. The formation of carbamoyl phosphate in the mitochondrial matrix is strongly stimulated by the allosteric effector N-acetylglutamate (see Fig. 17–14). Note that the terminal phosphate groups of two molecules of ATP are used to form one molecule of carbamoyl phosphate: two activation steps occur in the carbamoyl phosphate synthetase I reaction.
As we noted in Chapter 14, enzymes of many metabolic pathways are not randomly distributed within cellular compartments, but instead are clustered (p. 414). The product of one enzyme is often channeled directly to the next enzyme in the pathway. In the urea cycle, mitochondrial and cytosolic enzymes appear to be clustered in this way. The citrulline transported out of the mitochondria is not diluted into the general pool of metabolites in the cytosol. Instead, each molecule of citrulline is passed directly into the active site of a molecule of argininosuccinate synthetase. This channeling continues for argininosuccinate, arginine, and ornithine. Only the urea is released into the general pool within the cytosol.
Figure 17–13  The “Krebs bicycle”, composed of the urea cycle on the right, which meshes with the aspartate-argininosuccinate shunt of the citric acid cycle on the left. Fumarate produced in the cytosol by argininosuccinate lyase of the urea cycle enters the citric acid cycle in the mitochondrion and is converted in several steps to oxaloacetate. Oxaloacetate accepts an amino group from glutamate by transamination, and the aspartate thus formed leaves the mitochondrion and donates its amino group to the urea cycle in the argininosuccinate synthetase reaction. Intermediates in the citric acid cycle are boxed.

The fumarate produced in the argininosuccinate lyase reaction is also an intermediate of the citric acid cycle. Fumarate enters the mitochondria, where the combined activities of fumarase (fumarate hydratase) (p. 458) and malate dehydrogenase (p. 459) transform fumarate into oxaloacetate (Fig. 17–13). Aspartate, which acts as a nitrogen donor in the urea cycle reaction catalyzed by argininosuccinate synthetase in the cytosol, is formed from oxaloacetate by transamination from glutamate; the other product of this transamination is α-ketoglutarate, another intermediate of the citric acid cycle. Because the reactions of the urea and citric acid cycles are inextricably intertwined, together they have been called the “Krebs bicycle”.

Figure 17–14  Synthesis of N-acetylglutamate, the allosteric activator of carbamoyl phosphate synthetase I, is stimulated by high concentrations of arginine. Increasing arginine levels signal the need for more flux through the urea cycle.

The flux of nitrogen through the urea cycle varies with the composition of the diet. When the diet is primarily protein, the use of the carbon skeletons of amino acids for fuel results in the production of much urea from the excess amino groups. During severe starvation, when breakdown of muscle protein supplies much of the metabolic fuel, urea production also increases substantially, for the same reason.

These changes in demand for urea cycle activity are met in the long term by regulation of the rates of synthesis of the urea cycle enzymes and carbamoyl phosphate synthetase I in the liver. All five enzymes are synthesized at higher rates during starvation or in animals on very high-protein diets than in well-fed animals on diets containing primarily carbohydrates and fats. Animals on protein-free diets produce even lower levels of the urea cycle enzymes.

On a shorter time scale, allosteric regulation of at least one key enzyme is involved in adjusting flux through the cycle. The first enzyme in the pathway, carbamoyl phosphate synthetase I, is allosterically activated by N-acetylglutamate, which is synthesized from acetyl-CoA and glutamate (Fig. 17–14). N-Acetylglutamate synthase is in turn activated by arginine, a urea cycle intermediate that accumulates when urea production is too slow to accommodate the ammonia produced by amino acid catabolism.

The urea cycle brings together two amino groups and HCO3 to form a molecule of urea, which diffuses from the liver into the bloodstream, thence to be excreted into the urine by the kidneys. The overall equation of the urea cycle is

2NH4+ + HCO3 + 3ATP4− + H2O   →   urea + 2ADP3− + 4Pi2− + AMP2− + 5H+

The synthesis of one molecule of urea requires four high-energy phosphate groups. Two ATPs are required to make carbamoyl phosphate, and one ATP is required to make argininosuccinate. In the latter reaction, however, the ATP undergoes a pyrophosphate cleavage to AMP and pyrophosphate, which may be hydrolyzed to yield two Pi.

It has been estimated that, because of the necessity of excreting nitrogen as urea instead of ammonia, ureotelic animals lose about 15% of the energy of the amino acids from which the urea was derived. This loss is counterbalanced by metabolic adaptations in some ruminant animals. The cow transfers much urea from its blood into the rumen, where microorganisms use it as a source of ammonia to manufacture amino acids, which are then absorbed and used by the cow. Urea is sometimes added to cattle feed as an inexpensive nitrogen supplement. The recycling of urea not only reduces the net investment of chemical energy, it also reduces the requirements for protein intake and urine production. This can be important for ruminants that must subsist on a low-protein grass diet in a dry environment. The camel, by transferring urea into its gastrointestinal tract and recycling it like the cow, greatly reduces the water loss connected with the urinary excretion of urea. This is one of several biochemical and physiological adaptations that enables the camel to survive on a very limited water intake.

People with genetic defects in any enzyme involved in the formation of urea have an impaired ability to convert ammonia to urea. They cannot tolerate a protein-rich diet because amino acids ingested in excess of the minimum daily requirements for protein synthesis would be deaminated in the liver, producing free ammonia in the blood. As we have seen, ammonia is toxic and causes mental disorders, retarded development, and, in high amounts, coma and death. Humans, however, are incapable of synthesizing half of the 20 standard amino acids, and these essential amino acids (Table 17–1) must be provided in the diet.

Figure 17–15  The essential amino acids (those with carbon skeletons that cannot be synthesized by animals and must be obtained in the diet) can be synthesized from the corresponding α-keto acids by transamination. The dietary requirement for essential amino acids can therefore be met by the α-keto acid skeletons. RE and RN represent R groups of essential and nonessential amino acids, respectively.
Patients with defects in the urea cycle are often treated by substituting in the diet the α-keto acid analogs of the essential amino acids, which are the indispensable parts of the amino acids. The α-keto acid analogs can then accept amino groups from excess nonessential amino acids by aminotransferase action (Fig. 17–15). In this way the essential amino acids are made available for biosynthesis, and nonessential amino acids are kept from delivering their amino groups to the blood in the form of ammonia.
Figure 17–16  A view on San Lorenzo Island, one of the guano islands off the coast of Peru. Hundreds of thousands of “gooney” birds nest on these islands, and over the centuries, enormous clifflike deposits of guano, which are largely solid uric acid, have built up. Guano is a valuable fertilizer because of its nitrogen content.

Urea synthesis is not the only, or even the most common, pathway among organisms for excreting ammonia. The basis for differences in the molecular form in which amino groups are excreted lies in the anatomy and physiology of different organisms in relation to their usual habitat. Bacteria and free-living protozoa simply release ammonia to their aqueous environment, in which it is diluted and thus made harmless. In the bony fishes (ammonotelic animals), ammonia is rapidly cleared from the blood at the gills by the large volume of water passing through these respiratory structures. Although quite sensitive to NH3, fish are relatively tolerant of NH4+ . Liver is also the primary site of amino acid catabolism in fish, and NH4+ produced by transdeamination is simply released from the liver into the blood for transport to the gills and excretion. The bony fishes thus do not require a complex urinary system to excrete ammonia.

Organisms that excrete ammonia could not survive in an environment in which water is limited. The evolution of terrestrial species depended upon mutations that conferred the ability to convert ammonia to nontoxic substances that could be excreted in a small volume of water. Two main methods of excreting nitrogen have evolved: conversion to either urea or uric acid.

The importance of the habitat in excretion of amino nitrogen is illustrated by the change in the pathway of nitrogen excretion as the tadpole undergoes metamorphosis into the adult frog. Tadpoles are entirely aquatic and excrete amino nitrogen as ammonia through their gills. The tadpole liver lacks the necessary enzymes to make urea, but during metamorphosis it begins synthesizing these enzymes and loses the ability to excrete ammonia. In the adult frog, which is more terrestrial in habit, amino nitrogen is excreted almost entirely as urea.

In birds and reptiles, availability of water is an especially important consideration. Excretion of urea into urine requires simultaneous excretion of a relatively large volume of water; the weight of the required water would impede flight in birds, and reptiles living in arid environments must conserve water. Instead, these animals convert

amino nitrogen into uric acid (Fig. 17–4), a relatively insoluble compound that is extracted as a semisolid mass of uric acid crystals with the feces. For the advantage of excreting amino nitrogen in the form of solid uric acid, birds and reptiles must carry out considerable metabolic work; uric acid is a purine (see Fig. 12–1), and the biosynthesis of uric acid is a complex energy-requiring process that is part of the purine catabolic pathway (Chapter 21).

On many islands off the coast of South America, which serve as immense rookeries for sea birds, uric acid is deposited in enormous amounts (Fig. 17–16). These huge guano deposits are used as fertilizer, thus returning organic nitrogen to the soil, to be used again for the synthesis of amino acids by plants and soil microorganisms (Chapter 21).
Figure 17–17  A summary of the points of entry of the standard amino acids into the citric acid cycle. (The boxes around the amino acids are color-matched to the end-products (shaded) of the catabolic pathways, here and in figures throughout the rest of this chapter.) Some amino acids are listed more than once; these are broken down to yield different fragments, each of which enters the citric acid cycle at a different point. This scheme represents the major catabolic pathways in vertebrate animals, but there are minor variations from organism to organism. Threonine, for instance, is degraded into acetyl-CoA via pyruvate in some organisms via a pathway illustrated in Fig. 17–22.

There are 20 standard amino acids in proteins, with a variety of carbon skeletons. Correspondingly, there are 20 different catabolic pathways for amino acid degradation. In humans, these pathways taken together normally account for only 10 to 15% of the body's energy production. Therefore, the individual amino acid degradative pathways are not nearly as active as glycolysis and fatty acid oxidation. In addition, the activity of the catabolic pathways can vary greatly from one amino acid to the next, depending upon the balance between requirements for biosynthetic processes and the amounts of a given amino acid available. For this reason, we shall not examine them all in detail. The 20 catabolic pathways converge to form only five products, all of which enter the citric acid cycle. From here the carbons can be diverted to gluconeogenesis or ketogenesis, or they can be completely oxidized to CO2 and H2O (Fig. 17–17).

All or part of the carbon skeletons of ten of the amino acids are ultimately broken down to yield acetyl-CoA. Five amino acids are converted into α-ketoglutarate, four into succinyl-CoA, two into fumarate, and two into oxaloacetate. The individual pathways for the 20 amino acids will be summarized by means of flow diagrams, each leading to a specific point of entry into the citric acid cycle. In these diagrams the amino acid carbon atoms that enter the citric acid cycle are shown in color. Note that some amino acids appear more than once, reflecting the fact that different parts of their carbon skeletons have different fates. Some of the enzymatic reactions in these pathways that are particularly noteworthy for their mechanisms or their medical significance will be singled out for special discussion.

A variety of interesting chemical rearrangements are found among the amino acid catabolic pathways. Before examining the pathways themselves, it is useful to note classes of reactions that recur and to introduce the enzymatic cofactors required. We have already considered one important class, the transamination reactions requiring pyridoxal phosphate. Another common type of reaction in amino acid catabolism is a one-carbon transfer. One-carbon transfers usually involve one of three cofactors: biotin, tetrahydrofolate, or S-adenosylmethionine (Fig. 17–18). These cofactors are used to transfer one-carbon groups in different

oxidation states. The most oxidized state of carbon, CO2, is transferred by biotin (see Fig. 15–13). The remaining two cofactors are especially important in amino acid and nucleotide metabolism. Tetrahydrofolate is generally involved in transfers of one-carbon groups in the intermediate oxidation states, and S-adenosylmethionine in transfers of methyl groups, the most reduced state of carbon.

Tetrahydrofolate (H4 folate) consists of a substituted pteridine, p-aminobenzoate, and glutamate linked together as in Figure 17–18. This cofactor is synthesized in bacteria and its precursor, folate, is a

Figure 17–18  The structures of enzyme cofactors important
in one-carbon transfer reactions. The nitrogen atoms to which
one-carbon groups are attached in tetrahydrofolate are shown
in blue.
vitamin for mammals. The one-carbon group, in any of three oxidation states, is bonded to N-5 or N-10 or to both (Fig. 17–19). The most reduced form of the cofactor carries a methyl group, a more oxidized form carries a methylene group, and the most oxidized forms carry a methenyl, formyl, or formimino group. The different forms of tetrahydrofolate are interconvertible and serve as donors of one-carbon units in a variety of biosynthetic reactions. The major source of one-carbon units for tetrahydrofolate is the carbon removed in the conversion of serine to glycine, producing N5,N10-methylenetetrahydrofolate (Fig. 17–19).

Figure 17–19  Conversions of one-carbon units on tetrahydrofolate.
The different forms are grouped according to oxidation states, with the
most reduced at the top and most oxidized at the bottom. All of the forms
shown within a single shaded box are at the same oxidation state.
The enzymatic transfer of formyl groups (as in purine synthesis (Fig. 21–27)
and formation of formylmethionine in prokaryotes (p. 917)) generally
uses N10-formyltetrahydrofolate rather than N5-formyltetrahydrofolate.
The latter species is significantly more stable, and hence is not as good
a donor of formyl groups. Over time, the equilibria of the reactions that
interconnect these species will favor formation of N5-formyltetrahydrofolate.
The N5 species must be converted to N10-formyltetrahydrofolate in a reaction
that requires ATP because of its unfavorable equilibrium. Little is known
about the mechanism of this reaction. Note that N5-formiminotetrahydrofolate
is derived from histidine in a pathway shown in Fig. 17–29.

Although tetrahydrofolate can carry a methyl group at N-5, the methyl group’s transfer potential is insufficient for most biosynthetic reactions. S-Adenosylmethionine (adoMet) is more commonly used for methyl group transfers. It is synthesized from ATP and methionine

by the action of methionine adenosyl transferase (Fig. 17–20). This reaction is unusual in that the nucleophilic sulfur atom of methionine attacks at the 5′ carbon of the ribose moiety of ATP, releasing triphosphate, rather than attacking at one of the phosphorus atoms. The triphosphate is cleaved to Pi and PPi on the enzyme, and the PPi is later cleaved by inorganic pyrophosphatase, so that three bonds, two of which are high-energy bonds, are broken in this reaction. The only other reaction known in which triphosphate is displaced from ATP occurs in the synthesis of coenzyme B12 (see Box 16–2, Fig. 3).

Figure 17–20  Synthesis of methionine and S-adenosylmethionine as part of an
activated methyl cycle. The methyl group donor in the methionine synthase
reaction is methylcobalamin in some organisms. S-Adenosylmethionine, which has
a positively charged sulfur (and is thus a sulfonium ion), is a powerful methylating
agent in a number of biosynthetic reactions. The methyl group acceptor is
designated R.

S-Adenosylmethionine is a potent alkylating agent by virtue of its destabilizing sulfonium ion. The methyl group is subject to attack by nucleophiles and is about 1,000 times more reactive than the methyl group of N5-methyltetrahydrofolate.

Transfer of a methyl group from S-adenosylmethionine to an acceptor yields S-adenosylhomocysteine, which is subsequently broken down to homocysteine and adenosine (Fig. 17–20). Methionine is regenerated by the transfer of a methyl group to homocysteine in a reaction catalyzed by methionine synthase. One form of this enzyme is common in bacteria and uses N5-methyltetrahydrofolate as a methyl donor. Another form that occurs in bacteria and mammals uses methylcobalamin derived from coenzyme B12. This reaction and the rearrangement of L-methylmalonyl-CoA to succinyl-CoA (Box 16–2, Fig. 1a) are the only coenzyme B12-dependent reactions known in mammals. Methionine is reconverted to S-adenosylmethionine to complete an activated methyl cycle (Fig. 17–20).

Tetrahydrobiopterin is another cofactor introduced in these pathways, but it is not involved in one-carbon transfers. Tetrahydrobiopterin is structurally related to the flavin coenzymes, and it participates in biological oxidation reactions. It belongs to a widespread class of biological compounds called pterins (Fig. 17–21), and we will consider its mode of action when we discuss phenylalanine degradation.

Figure 17–21  Tetrahydrobiopterin and its parent compound, pterin.
Tetrahydrobiopterin is a cofactor for the enzyme phenylalanine hydroxylase.

The carbon skeletons of ten amino acids yield acetyl-CoA, which enters the citric acid cycle directly (Fig. 17–17). Five of the ten are degraded to acetyl-CoA via pyruvate. The other five are converted into acetyl-CoA and/or acetoacetyl-CoA, which is then cleaved to form acetyl-CoA.

The five amino acids entering via pyruvate are alanine, glycine, serine, cysteine, and tryptophan (Fig. 17–22). In some organisms threonine is also degraded to form acetyl-CoA, as shown in Figure 17–22; in humans it is degraded to succinyl-CoA, as described later. Alanine yields pyruvate directly on transamination with α-ketoglutarate, and the side chain of tryptophan is cleaved to yield alanine and thus pyruvate. Cysteine is converted to pyruvate in two steps, one to remove the sulfur atom, the other a transamination. Serine is converted to pyruvate by serine dehydratase. Both the β-hydroxyl and the α-amino groups of serine are removed in this single PLP-dependent reaction (an analogous reaction with threonine is shown in Fig. 17–30). Glycine has two pathways. It can be converted into serine by enzymatic addition of a hydroxymethyl group (Fig. 17–23a). This reaction, catalyzed by serine hydroxymethyl transferase, requires the coenzymes tetrahydrofolate and pyridoxal phosphate. The second pathway for glycine, which predominates in animals, involves its oxidative cleavage into CO2, NH4+, and a methylene group (–CH2–) (Fig. 17–23b). This readily reversible reaction, catalyzed by glycine synthase, also requires tetrahydrofolate, which accepts the methylene group. In this oxidative cleavage pathway the two carbon atoms of glycine do not enter the citric acid cycle. One is lost as CO2, and the other becomes the methylene group of N5,N10-methylene-tetrahydrofolate (Fig. 17–19), which is used as a one-carbon group donor in certain biosynthetic pathways.

Figure 17–22  Outline of the catabolic pathways for alanine, glycine, serine, cysteine, tryptophan, and threonine. The fate of the indole group of tryptophan is shown in Fig. 17–24. Details of the glycine-to-serine conversion, and a second fate for glycine, are shown in Fig. 17–23. In some organisms (including humans) threonine is degraded to succinyl-CoA by another pathway (Fig. 17–30). There are several pathways for cysteine degradation, all of which lead to pyruvate. The enzyme serine hydroxymethyl transferase contains both pyridoxal phosphate and tetrahydrofolate. The threonine cleavage reaction shown here is catalyzed by the same enzyme. Carbon atoms here and in subsequent figures are color-coded as necessary to trace their fates, in addition to the color-coding for pathways described in Fig. 17–17.

Figure 17–23  Two metabolic fates of glycine: (a) conversion to serine and (b) breakdown to CO2 and ammonia. The cofactor tetrahydrofolate carries one-carbon units in both of these reactions. The structure of H4 folate is shown in Fig. 17–18, and its role as a cofactor in one-carbon transfers in Fig. 17–19.
Portions of the carbon skeleton of six amino acids – tryptophan, lysine, phenylalanine, tyrosine, leucine, and isoleucine – yield acetyl-CoA and/or acetoacetyl-CoA; the latter is then converted into acetyl-CoA (Fig. 17–24). Some of the final steps in the degradative pathways for leucine, lysine, and tryptophan resemble steps in the oxidation of fatty acids. The breakdown of two of these six amino acids deserves special mention.

Figure 17–24  Summary of the catabolic fates of tryptophan, lysine, phenylalanine, tyrosine, leucine, and isoleucine, which donate some of their carbons (those in red) to acetyl-CoA. Tryptophan, phenylalanine, tyrosine, and isoleucine also contribute carbons (in blue) as pyruvate or citric acid cycle intermediates. The phenylalanine pathway is described in more detail in Fig. 17–26. The fate of nitrogen atoms is not traced in this scheme. In most cases they are transferred to α-ketoglutarate to form glutamate.
The dehydration of tryptophan is the most complex of all the pathways of amino acid catabolism in animal tissues; portions of tryptophan (six carbons total) yield acetyl-CoA by two different pathways, one via pyruvate and one via acetoacetyl-CoA. Some of the intermediates in tryptophan catabolism are required precursors for biosynthesis of other important biomolecules (Fig. 17–25), including nicotinate, a precursor of NAD and NADP. In plants, the growth factor indoleacetate is derived from tryptophan by an oxidative pathway. Tryptophan is also the parent, by a different pathway, of the neurotransmitter serotonin. Some of these biosynthetic pathways are described in more detail in Chapter 21.

Figure 17–25  The aromatic rings of tryptophan are precursors of nicotinate, indoleacetate, and serotonin. Colored atoms are used to trace the source of the ring atoms in nicotinate.

The breakdown of phenylalanine is noteworthy because genetic defects in the enzymes of phenylalanine catabolism lead to several dif ferent inheritable human diseases (Fig. 17–26), as discussed below. Phenylalanine and its oxidation product tyrosine are degraded into two fragments, each of which can enter the citric acid cycle, but at different points. Four of the nine carbon atoms of phenylalanine and tyrosine yield free acetoacetate, which is converted into acetoacetyl-CoA. A second four-carbon fragment of tyrosine and phenylalanine is recovered as fumarate. Eight of the nine carbon atoms of these two amino acids thus enter the citric acid cycle; the remaining carbon is lost as CO2. Phenylalanine, after its hydroxylation to yield tyrosine, is also the precursor of the hormones epinephrine and norepinephrine, secreted by the adrenal medulla, the neurotransmitter dopamine, and melanin, the black pigment of skin and hair (Chapter 21).

Many different genetic defects in amino acid metabolism have been identified in humans (Table 17–2, p. 530). Most such defects cause specific intermediates to accumulate, a condition that can cause defective neural development and mental retardation.

The first enzyme in the catabolic pathway for phenylalanine (Fig. 17–26), phenylalanine hydroxylase, catalyzes the hydroxylation of phenylalanine to tyrosine. A genetic defect in phenylalanine hydroxylase is responsible for the disease phenylketonuria (PKU). Phenylketonuria is the most common cause of elevated levels of phenylalanine (hyperphenylalaninemia). Phenylalanine hydroxylase inserts one of the two oxygen atoms of O2 into phenylalanine to form the hydroxyl group of tyrosine; the other oxygen atom is reduced to H2O by the NADH also required in the reaction. This is one of a general class of reactions catalyzed by enzymes called mixed-function oxidases (see Box 20–1), all of which catalyze simultaneous hydroxylation of a substrate by O2 and reduction of the other oxygen atom of O2 to H2O. Phenylalanine hydroxylase requires a cofactor, tetrahydrobiopterin, which carries electrons from NADH to O2 in the hydroxylation of phenylalanine. During the hydroxylation reaction the coenzyme is oxidized to dihydrobiopterin (Fig. 17–27). It is subsequently reduced again by the enzyme dihydrobiopterin reductase in a reaction that requires NADH.

Figure 17–26  The normal pathway for conversion of phenylalanine and tyrosine into acetoacetyl-CoA and fumarate in humans. Genetic defects in each of the first four enzymes in this pathway are known to cause inheritable human diseases (shaded in red).
Chapter 18


Part X
Chapter G

E0:  See standard reduction potential.

E. coli (Escherichia coli):  A common bacterium found in the small intestine of vertebrates; the most well-studied organism.

electrochemical gradient:  The sum of the gradients of concentration and of electric charge of an ion across a membrane; the driving force for oxidative phosphorylation and photophosphorylation.

electrochemical potential:  The energy required to maintain a separation of charge and of concentration across a membrane.

electrogenic:  Contributing to an electrical potential across a membrane.

electron acceptor:  A substance that receives electrons in an oxidation–reduction reaction.

electron carrier:  A protein, such as a flavoprotein or a cytochrome, that can reversibly gain and lose electrons; functions in the transfer of electrons from organic nutrients to oxygen or some other terminal acceptor.

electron donor:  A substance that donates electrons in an oxidation–reduction reaction.

electron transfer:  Movement of electrons from substrates to oxygen via the carriers of the respiratory (electron transfer) chain.

electrophile:  An electron-deficient group with a strong tendency to accept electrons from an electron-rich group (nucleophile).

electrophoresis:  Movement of charged solutes in response to an electrical field; often used to separate mixtures of ions, proteins, or nucleic acids.

elongation factors:  Specific proteins required in the elongation of polypeptide chains by ribosomes.

eluate:  The effluent from a chromatographic column.

enantiomers:  Stereoisomers that are nonsuperimposable mirror images of each other.

end-product inhibition:  See feedback inhibition.

endergonic reaction:  A chemical reaction that consumes energy (that is, for which ΔG is positive).

endocrine glands:  Groups of cells specialized to synthesize hormones and secrete them into the blood to regulate other types of cells.

endocytosis:  The uptake of extracellular material by its inclusion within a vesicle (endosome) formed by an invagination of the plasma membrane.

endonuclease:  An enzyme that hydrolyzes the interior phosphodiester bonds of a nucleic acid; that is, it acts at points other than the terminal bonds.

endoplasmic reticulum:  An extensive system of double membranes in the cytoplasm of eukaryotic cells; it encloses secretory channels and is often studded with ribosomes (rough endoplasmic reticulum).

endothermic reaction:  A chemical reaction that takes up heat (that is, for which ΔH is positive).

energy charge:  The fractional degree to which the ATP/ADP/AMP system is filled with high-energy phosphate groups.

energy coupling:  The transfer of energy from one process to another.

enhancers:  DNA sequences that facilitate the expression of a given gene; may be located a few hundred, or even thousand, base pairs away from the gene.

enthalpy (H):  The heat content of a system.

enthalpy change (ΔH):  For a reaction, is approximately equal to the difference between the energy used to break bonds and the energy gained by the formation of new ones.

entropy (S):  The extent of randomness or disorder in a system.

enzyme:  A biomolecule, either protein or RNA, that catalyzes a specific chemical reaction. It does not affect the equilibrium of the catalyzed reaction; it enhances the rate of a reaction by providing a reaction path with a lower activation energy.

epimerases:  Enzymes that catalyze the reversible interconversion of two epimers.

epimers:  Two stereoisomers differing in configuration at one asymmetric center, in a compound having two or more asymmetric centers.

epithelial cell:  Any cell that forms part of the outer covering of an organism or organ.

epitope:  An antigenic determinant; the particular chemical group or groups within a macromolecule (antigen) to which a given antibody binds.

equilibrium:  The state of a system in which no further net change is occurring; the free energy is at a minimum.

equilibrium constant (Keq):  A constant, characteristic for each chemical reaction; relates the specific concentrations of all reactants and products at equilibrium at a given temperature and pressure.

erythrocyte:  A cell containing large amounts of hemoglobin and specialized for oxygen transport; a red blood cell.

Escherichia coli:  See E. coli.

essential amino acids:  Amino acids that cannot be synthesized by humans (and other vertebrates) and must be obtained from the diet.

essential fatty acids:  The group of polyunsaturated fatty acids produced by plants, but not by humans; required in the human diet.

ethanol fermentation:  See alcohol fermentation.

eukaryote:  A unicellular or multicellular organism with cells having a membrane-bounded nucleus, multiple chromosomes, and internal organelles.

excited state:  An energy-rich state of an atom or molecule; produced by the absorption of light energy.

exergonic reaction:  A chemical reaction that proceeds with the release of free energy (that is, for which ΔG is negative).

exocytosis:  The fusion of an intracellular vesicle with the plasma membrane, releasing the vesicle contents to the extracellular space.

exon:  The segment of a eukaryotic gene that encodes a portion of the final product of the gene; a portion that remains after posttranscriptional processing and is transcribed into a protein or incorporated into the structure of an RNA. See intron.

exonuclease:  An enzyme that hydrolyzes only those phosphodiester bonds that are in the terminal positions of a nucleic acid.

exothermic reaction:  A chemical reaction that releases heat (that is, for which ΔH is negative).

expression vector:  See vector.

facilitated diffusion:  Diffusion of a polar substance across a biological membrane through a protein transporter; also called passive diffusion or passive transport.

facultative cells:  Cells that can live in the presence or absence of oxygen.

FAD (flavin adenine dinucleotide):  The coenzyme of some oxidation–reduction enzymes; it contains riboflavin.

fatty acid:  A long-chain aliphatic carboxylic acid found in natural fats and oils; also a component of membrane phospholipids and glycolipids.

feedback inhibition:  Inhibition of an allosteric enzyme at the beginning of a metabolic sequence by the end product of the sequence; also known as end-product inhibition.

fermentation:  Energy-yielding anaerobic breakdown of a nutrient molecule, such as glucose, without net oxidation; yields lactate, ethanol, or some other simple product.

fibroblast:  A cell of the connective tissue that secretes connective tissue proteins such as collagen.

fibrous proteins:  Insoluble proteins that serve in a protective or structural role; contain polypeptide chains that generally share a common secondary structure.

fingerprinting:  See peptide mapping.

first law of thermodynamics:  The law stating that in all processes, the total energy of the universe remains constant.

Fischer projection formulas:  See projection formulas.

5′ end:  The end of a nucleic acid that lacks a nucleotide bound at the 5′ position of the terminal residue.

flagellum:  A cell appendage used in propulsion. Bacterial flagella have a much simpler structure than eukaryotic flagella, which are similar to cilia.

flavin-linked dehydrogenases:  Dehydrogenases requiring one of the riboflavin coenzymes, FMN or FAD.

flavin nucleotides:  Nucleotide coenzymes (FMN and FAD) containing riboflavin.

flavoprotein:  An enzyme containing a flavin nucleotide as a tightly bound prosthetic group.

fluid mosaic model:  A model describing biological membranes as a fluid lipid bilayer with embedded proteins; the bilayer exhibits both structural and functional asymmetry.

fluorescence:  Emission of light by excited molecules as they revert to the ground state.

FMN (flavin mononucleotide):  Riboflavin phosphate, a coenzyme of certain oxidation–reduction enzymes.

footprinting:  A technique for identifying the nucleic acid sequence bound by a DNA- or RNA-binding protein.

frame shift:  A mutation caused by insertion or deletion of one or more paired nucleotides, changing the reading frame of codons during protein synthesis; the polypeptide product has a garbled amino acid sequence beginning at the mutated codon.

free energy (G):  The component of the total energy of a system that can do work at constant temperature and pressure.

free energy of activation (ΔG):  See activation energy.

free-energy change (ΔG):  The amount of free energy released (negative ΔG) or absorbed (positive ΔG) in a reaction at constant temperature and pressure.

free radical:  See radical.

functional group:  The specific atom or group of atoms that confers a particular chemical property on a biomolecule.

furanose:  A simple sugar containing the five-membered furan ring.

fusion protein:  (1) A family of proteins that facilitate membrane fusion. (2) The protein product of a gene created by the fusion of two distinct genes.

futile cycle:  A set of enzyme-catalyzed cyclic reactions that results in release of thermal energy by the hydrolysis of ATP.

ΔG°’:  See standard free-energy change.

gametes:  Reproductive cells with a haploid gene content; sperm or egg cells.

gangliosides:  Sphingolipids, containing complex oligosaccharides as head groups; especially common in nervous tissue.

gel filtration:  A chromatographic procedure for the separation of a mixture of molecules on the basis of size; based on the capacity of porous polymers to exclude solutes above a certain size.

gene:  A chromosomal segment that codes for a single functional polypeptide chain or RNA molecule.

gene expression:  Transcription and, in the case of proteins, translation to yield the product of a gene; a gene is expressed when its biological product is present and active.

gene splicing:  The enzymatic attachment of one gene, or part of a gene, to another.

general acid–base catalysis:  Catalysis involving proton transfer(s) to or from a molecule other than water.

genetic code:  The set of triplet code words in DNA (or mRNA) coding for the amino acids of proteins.

genetic information:  The hereditary information contained in a sequence of nucleotide bases in chromosomal DNA or RNA.

genetic map:  A diagram showing the relative sequence and position of specific genes along a chromosome.

genome:  All the genetic information encoded in a cell or virus.

genotype:  The genetic constitution of an organism, as distinct from its physical characteristics, or phenotype.

geometric isomers:  Isomers related by rotation about a double bond; also called cis and trans isomers.

germ-line cell:  A type of animal cell that is formed early in embryogenesis and may multiply by mitosis or may produce, by meiosis, cells that develop into gametes (egg or sperm cells).

globular proteins:  Soluble proteins with a globular (somewhat rounded) shape.

glucogenic amino acids:  Amino acids with carbon chains that can be metabolically converted into glucose or glycogen via gluconeogenesis.

gluconeogenesis:  The biosynthesis of a carbohydrate from simpler, noncarbohydrate precursors such as oxaloacetate or pyruvate.

glycan:  Another term for polysaccharide; a polymer of monosaccharide units joined by glycosidic bonds.

glycerophospholipid:  An amphipathic lipid with a glycerol backbone; fatty acids are ester-linked to C-1 and C-2 of glycerol, and a polar alcohol is attached through a phosphodiester linkage to C-3.

glycolipid:  A lipid containing a carbohydrate group.

glycolysis:  The catabolic pathway by which a molecule of glucose is broken down into two molecules of pyruvate.

glycoprotein:  A protein containing a carbohydrate group.

glycosaminoglycan:  A heteropolysaccharide of two alternating units: one is either N-acetylglucosamine or N-acetylgalactosamine; the other is a uronic acid (usually glucuronic acid). Formerly called mucopolysaccharide.

glycosidic bonds:  Bonds between a sugar and another molecule (typically an alcohol, purine, pyrimidine, or sugar) through an intervening oxygen or nitrogen atom; the bonds are classified as O-glycosidic or N-glycosidic, respectively.

glyoxylate cycle:  A variant of the citric acid cycle, for the net conversion of acetate into succinate and, eventually, new carbohydrate; present in bacteria and some plant cells.

glyoxysome:  A specialized peroxisome containing the enzymes of the glyoxylate cycle; found in cells of germinating seeds.

Golgi complex:  A complex membranous organelle of eukaryotic cells; functions in the posttranslational modification of proteins and their secretion from the cell or incorporation into the plasma membrane or organellar membranes.

gram molecular weight:  The weight in grams of a compound that is numerically equal to its molecular weight; the weight of 1 mole.

grana:  Stacks of thylakoids, flattened membranous sacs or discs, in chloroplasts.

ground state:  The normal, stable form of an atom or molecule; as distinct from the excited state.

group transfer potential:  A measure of the ability of a compound to donate an activated group (such as a phosphate or acyl group); generally expressed as the standard free energy of hydrolysis.

half-life:  The time required for the disappearance or decay of one-half of a given component in a system.

haploid:  Having a single set of genetic information; describing a cell with one chromosome of each type.

Haworth perspective formulas:  A method for representing cyclic chemical structures so as to define the configuration of each substituent group; the method commonly used for representing sugars.

helicase:  An enzyme that catalyzes the separation of strands in a DNA molecule before replication.

helix, α:  See α helix.

heme:  The iron-porphyrin prosthetic group of heme proteins.

heme protein:  A protein containing a heme as a prosthetic group.

hemoglobin:  A heme protein in erythrocytes; functions in oxygen transport.

Henderson–Hasselbalch equation:  An equation relating the pH, the pKa, and the ratio of the concentrations of the proton-acceptor (A) and proton-donor (HA) species in a solution.

hepatocyte:  The major cell type of liver tissue.

heteroduplex DNA:  Duplex DNA containing complementary strands derived from two different DNA molecules with similar sequences, often as a product of genetic recombination.

heteropolysaccharide:  A polysaccharide containing more than one type of sugar.

heterotroph:  An organism that requires complex nutrient molecules, such as glucose, as a source of energy and carbon.

heterotropic enzyme:  An allosteric enzyme requiring a modulator other than its substrate.

hexose:  A simple sugar with a backbone containing six carbon atoms.

high-energy compound:  A compound that on hydrolysis undergoes a large decrease in free energy under standard conditions.

high-performance liquid chromatography (HPLC):  Chromatographic procedures, often conducted at relatively high pressures, using automated equipment that permits refined and highly reproducible profiles.

Hill reaction:  The evolution of oxygen and the photoreduction of an artificial electron acceptor by a chloroplast preparation in the absence of carbon dioxide.

histones:  The family of five basic proteins that associate tightly with DNA in the chromosomes of all eukaryotic cells.

Holliday intermediate:  An intermediate in genetic recombination in which two double-stranded DNA molecules are joined by virtue of a reciprocal crossover involving one strand of each molecule.

holoenzyme:  A catalytically active enzyme including all necessary subunits, prosthetic groups, and cofactors.

homeobox:  A conserved DNA sequence of 180 base pairs encoding a protein domain found in many proteins that play a regulatory role in development.

homeodomain:  The protein domain encoded by the homeobox.

homeostasis:  The maintenance of a dynamic steady state by regulatory mechanisms that compensate for changes in external circumstances.

homeotic genes:  Genes that regulate the development of the pattern of segments in the Drosophila body plan; similar genes are found in most vertebrates.

homologous genetic recombination:  Recombination between two DNA molecules of similar sequence, occurring in all cells; occurs during meiosis and mitosis in eukaryotes.

homologous proteins:  Proteins having sequences and functions similar in difierent species; for example, the hemoglobins.

homopolysaccharide:  A polysaccharide made up of only one type of monosaccharide unit.

homotropic enzyme:  An allosteric enzyme that uses its substrate as a modulator.

hormone:  A chemical substance synthesized in small amounts by an endocrine tissue and carried in the blood to another tissue, where it acts as a messenger to regulate the function of the target tissue or organ.

hormone receptor:  A protein in, or on the surface of, target cells that binds a specific hormone and initiates the cellular response.

hydrogen bond:  A weak electrostatic attraction between one electronegative atom (such as oxygen or nitrogen) and a hydrogen atom covalently linked to a second electronegative atom.

hydrolases:  Enzymes (proteases, lipases, phosphatases, nucleases, for example) that catalyze hydrolysis reactions.

hydrolysis:  Cleavage of a bond, such as an anhydride or peptide bond, by the addition of the elements of water, yielding two or more products.

hydronium ion:  The hydrated hydrogen ion (H3O+).

hydropathy index:  A scale that expresses the relative hydrophobic and hydrophilic tendencies of a chemical group.

hydrophilic:  Polar or charged; describing molecules or groups that associate with (dissolve easily in) water.

hydrophobic:  Nonpolar; describing molecules or groups that are insoluble in water.

hydrophobic interactions:  The association of nonpolar groups, or compounds, with each other in aqueous systems, driven by the tendency of the surrounding water molecules to seek their most stable (disordered) state.

hyperchromic effect:  The large increase in light absorption at 260 nm occurring as a double-helical DNA is melted (unwound).

immune response:  The capacity of a vertebrate to generate antibodies to an antigen, a macromolecule foreign to the organism.

immunoglobulin:  An antibody protein generated against, and capable of binding specifically to, an antigen.

in vitro:  “In glass”; that is, in the test tube.

in vivo:  “In life”; that is, in the living cell or organism.

induced fit:  A change in the conformation of an enzyme in response to substrate binding that renders the enzyme catalytically active; also used to denote changes in the conformation of any macromolecule in response to ligand binding such that the binding site of the macromolecule better conforms to the shape of the ligand.

inducer:  A signal molecule that, when bound to a regulatory protein, produces an increase in the expression of a given gene.

induction:  An increase in the expression of a gene in response to a change in the activity of a regulatory protein.

informational macromolecules:  Biomolecules containing information in the form of specific sequences of different monomers; for example, many proteins, lipids, polysaccharides, and nucleic acids.

initiation codon:  AUG (sometimes GUG in prokaryotes); codes for the first amino acid in a polypeptide sequence: N-formylmethionine in prokaryotes, and methionine in eukaryotes.

initiation complex:  A complex of a ribosome with an mRNA and the initiating Met-tRNAMet or fMet-tRNAfMet, ready for the elongation steps.

inorganic pyrophosphatase:  An enzyme that hydrolyzes a molecule of inorganic pyrophosphate to yield two molecules of (ortho) phosphate; also known as pyrophosphatase.

insertion mutation:  A mutation caused by insertion of one or more extra bases, or a mutagen, between two successive bases in DNA.

insertion sequence:  Specific base sequences at either end of a transposable segment of DNA.

integral membrane proteins:  Proteins firmly bound to a membrane by hydrophobic interactions; as distinct from peripheral proteins.

intercalating mutagen:  A mutagen that inserts itself between two successive bases in a nucleic acid, causing a frame-shift mutation.

intercalation:  Insertion between two stacked aromatic or planar rings; for example, the insertion of a planar molecule between two successive bases in a nucleic acid.

interferons:  A class of glycoproteins with antiviral activities.

intermediary metabolism:  In cells, the enzyme-catalyzed reactions that extract chemical energy from nutrient molecules and utilize it to synthesize and assemble cell components.

intron (intervening sequence):  A sequence of nucleotides in a gene that is transcribed but excised before the gene is translated.

ion channel:  An integral membrane protein that provides for the regulated transport of a specific ion, or ions, across a membrane.

ion-exchange resin:  A polymeric resin that contains fixed charged groups; used in chromatographic columns to separate ionic compounds.

ion product of water (Kw):  The product of the concentrations of H+ and OH in pure water: Kw = [H+][OH] = 1 × 10−14 at 25 °C.

ionizing radiation:  A type of radiation, such as x rays, that causes loss of electrons from some organic molecules, thus making them more reactive.

ionophore:  A compound that binds one or more metal ions and is capable of diffusing across a membrane, carrying the bound ion.

iron–sulfur center:  A prosthetic group of certain redox proteins involved in electron transfers; Fe2+ or Fe3+ is bound to inorganic sulfur and to Cys groups in the protein.

isoelectric focusing:  An electrophoretic method for separating macromolecules on the basis of their isoelectric pH.

isoelectric pH (isoelectric point):  The pH at which a solute has no net electric charge and thus does not move in an electric field.

isoenzymes:  See isozymes.

isomerases:  Enzymes that catalyze the transformation of compounds into their positional isomers.

isomers:  Any two molecules with the same molecular formula but a different arrangement of molecular groups.

isoprene:  The hydrocarbon 2-methyl-1,3-butadiene, a recurring structural unit of the terpenoid biomolecules.

isothermal:  Occurring at constant temperature.

isotopes:  Stable or radioactive forms of an element that differ in atomic weight but are otherwise chemically identical to the naturally abundant form of the element; used as tracers.

isozymes:  Multiple forms of an enzyme that catalyze the same reaction but differ from each other in their amino acid sequence, substrate affinity, Vmax, and/or regulatory properties; also called isoenzymes.

keratins:  Insoluble protective or structural proteins consisting of parallel polypeptide chains in α-helical or β conformations.

ketogenic amino acids:  Amino acids with carbon skeletons that can serve as precursors of the ketone bodies.

ketone bodies:  Acetoacetate, D-β-hydroxybutyrate, and acetone; water-soluble fuels normally exported by the liver but overproduced during fasting or in untreated diabetes mellitus.

ketose:  A simple monosaccharide in which the carbonyl group is a ketone.

ketosis:  A condition in which the concentration of ketone bodies in the blood, tissues, and urine is abnormally high.

kinases:  Enzymes that catalyze the phosphorylation of certain molecules by ATP.

kinetics:  The study of reaction rates.

Krebs cycle:  See citric acid cycle.

lagging strand:  The DNA strand that, during replication, must be synthesized in the direction opposite to that in which the replication fork moves.

law of mass action:  The law stating that the rate of any given chemical reaction is proportional to the product of the activities (or concentrations) of the reactants.

leader:  A short sequence near the amino terminus of a protein or the 5′ end of an RNA that has a specialized targeting or regulatory function.

leading strand:  The DNA strand that, during replication, is synthesized in the same direction in which the replication fork moves.

leaky mutant:  A mutant gene that gives rise to a product with a detectable Ievel of biological activity.

leaving group:  The departing or displaced molecular group in a unimolecular elimination or a bimolecular substitution reaction.

lethal mutation:  A mutation that inactivates a biological function essential to the life of the cell or organism.

leucine zipper:  A protein structural motif involved in protein–protein interactions in many eukaryotic regulatory proteins; consists of two interacting α helices in which Leu residues in every seventh position are a prominent feature of the interacting surfaces.

leukotrienes:  A family of molecules derived from arachidonate; muscle contractants that constrict air passages in the lungs and are involved in asthma.

levorotatory isomer:  A stereoisomer that rotates the plane of plane-polarized light counterclockwise.

ligand:  A small molecule that binds specifically to a larger one; for example, a hormone is the ligand for its specific protein receptor.

light reactions:  The reactions of photosynthesis that require light and cannot occur in the dark; also known as the light-dependent reactions.

Lineweaver–Burk equation:  An algebraic transform of the Michaelis–Menten equation, allowing determination of Vmax and Km by extrapolation of [S] to infinity.

linking number:  The number of times one closed circular DNA strand is wound about another; the number of topological links holding the circles together.

lipases:  Enzymes that catalyze the hydrolysis of triacylglycerols.

lipid:  A small water-insoluble biomolecule generally containing fatty acids, sterols, or isoprenoid compounds.

lipoate (lipoic acid):  A vitamin for some microorganisms; an intermediate carrier of hydrogen atoms and acyl groups in α-keto acid dehydrogenases.

lipoprotein:  A lipid–protein aggregate that serves to carry water-insoluble lipids in the blood. The protein component alone is an apolipoprotein.

low-energy phosphate compound:  A phosphorylated compound with a relatively small standard free energy of hydrolysis.

lyases:  Enzymes that catalyze the removal of a group from a molecule to form a double bond, or the addition of a group to a double bond.

lymphocytes:  A subclass of leukocytes involved in the immune response. B lymphocytes synthesize and secrete antibodies; T lymphocytes either play a regulatory role in immunity or kill foreign and virus-infected cells.

lysis:  Destruction of a cell’s plasma membrane or of a bacterial cell wall, releasing the cellular contents and killing the cell.

lysogeny:  One of two outcomes of the infection of a host cell by a temperate phage. It occurs when the phage genome becomes repressed and is replicated as part of the host DNA; infrequently it may be induced, and the phage particles so produced cause the host cell to lyse.

lysosome:  A membrane-bounded organelle in the cytoplasm of eukaryotic cells; it contains many hydrolytic enzymes and serves as a degrading and recycling center for unneeded components.

macromolecule:  A molecule having a molecular weight in the range of a few thousand to many millions.

mass-action ratio:  For the reaction aA + dB ⇌ cC + dD, the ratio:

( [C]c [D]d ) / ( [A]a [B]b ).

matrix:  The aqueous contents of a cell or organelle (the mitochondrion, for example) with dissolved solutes.

meiosis:  A type of cell division in which diploid cells give rise to haploid cells destined to become gametes.

membrane transport:  Movement of a polar solute across a membrane via a specific membrane protein (a transporter).

messenger RNA (mRNA):  A class of RNA molecules, each of which is complementary to one strand of DNA; carries the genetic message from the chromosome to the ribosomes.

metabolism:  The entire set of enzyme-catalyzed transformations of organic molecules in living cells; the sum of anabolism and catabolism.

metabolite:  A chemical intermediate in the enzyme-catalyzed reactions of metabolism.

metalloprotein:  A protein having a metal ion as its prosthetic group.

metamerism:  Division of the body into segments; in insects, for example.

micelle:  An aggregate of amphipathic molecules in water, with the nonpolar portions in the interior and the polar portions at the exterior surface, exposed to water.

Michaelis–Menten constant (Km):  The substrate concentration at which an enzyme-catalyzed reaction proceeds at one-half its maximum velocity.

Michaelis–Menten equation:  The equation describing the hyperbolic dependence of the initial reaction velocity, V0, on substrate concentration, [S], in many enzyme-catalyzed reactions:

V0 = ( Vmax [S] ) / ( Km + [S] ).

Michaelis–Menten kinetics:  A kinetic pattern in which the initial rate of an enzyme-catalyzed reaction exhibits a hyperbolic dependence on substrate concentration.

microbodies:  Cytoplasmic, membrane-bounded vesicles containing peroxide-forming and peroxide-destroying enzymes; include lysosomes, peroxisomes, and glyoxysomes.

microfilaments:  Thin filaments composed of actin, found in the cytoplasm of eukaryotic cells; serve in structure and movement.

microsomes:  Membranous vesicles formed by fragmentation of the endoplasmic reticulum of eukaryotic cells; recovered by differential centrifugation.

microtubules:  Thin tubules assembled from two types of globular tubulin subunits; present in cilia, flagella, centrosomes, and other contractile or motile structures.

mitochondrion:  Membrane-bounded organelle in the cytoplasm of eukaryotes; contains the enzyme systems required for the citric acid cycle, fatty acid oxidation, electron transfer, and oxidative phosphorylation.

mitosis:  The multistep process in eukaryotic cells that results in the replication of chromosomes and cell division.

mixed-function oxidases (oxygenases):  Enzymes, often flavoproteins, that use molecular oxygen (O2) to simultaneously oxidize a substrate and a cosubstrate (commonly NADH or NADPH).

modulator:  A metabolite that, when bound to the allosteric site of an enzyme, alters its kinetic characteristics.

molar solution:  One mole of solute dissolved in water to give a total volume of 1,000 mL.

mole:  One gram molecular weight of a compound. See Avogadro’s number.

monoclonal antibodies:  Antibodies produced by a cloned hybridoma cell, which therefore are identical and directed against the same epitope of the antigen.

monolayer:  A single layer of oriented lipid molecules.

monoprotic acid:  An acid having only one dissociable proton.

monosaccharide:  A carbohydrate consisting of a single sugar unit.

mRNA:  See messenger RNA.

mucopolysaccharide:  An older name for a glycosaminoglycan.

multienzyme system:  A group of related enzymes participating in a given metabolic pathway.

mutarotation:  The change in specific rotation of a pyranose or furanose sugar or glycoside accompanying the equilibration of its α- and β-anomeric forms.

mutases:  Enzymes that catalyze the transposition of functional groups.

mutation:  An inheritable change in the nucleotide sequence of a chromosome.

myofibril:  A unit of thick and thin filaments of muscle fibers.

myosin:  A contractile protein; the major component of the thick filaments of muscle and other actin–myosin systems.

NAD, NADP (nicotinamide adenine dinucleotide, nicotinamide adenine dinucleotide phosphate):  Nicotinamide-containing coenzymes functioning as carriers of hydrogen atoms and electrons in some oxidation–reduction reactions.

native conformation:  The biologically active conformation of a macromolecule.

negative cooperativity:  A phenomenon of some multisubunit enzymes or proteins in which binding of a ligand or substrate to one subunit impairs binding to another subunit.

negative feedback:  Regulation of a biochemical pathway achieved when a reaction product inhibits an earlier step in the pathway.

neuron:  A cell of nervous tissue specialized for transmission of a nerve impulse.

neurotransmitter:  A low molecular weight compound (usually containing nitrogen) secreted from the terminal of a neuron and bound by a specific receptor in the next neuron; serves to transmit a nerve impulse.

nicotinamide adenine dinucleotide, nicotinamide adenine dinucleotide phosphate:  See NAD, NADP.

ninhydrin reaction:  A color reaction given by amino acids and peptides on heating with ninhydrin; widely used for their detection and estimation.

nitrogen cycle:  The cycling of various forms of biologically available nitrogen through the plant, animal, and microbial worlds, and through the atmosphere and geosphere.

nitrogen fixation:  Conversion of atmospheric nitrogen (N2) into a reduced, biologically available form by nitrogen-fixing organisms.

nitrogenase complex:  A system of enzymes capable of reducing atmospheric nitrogen to ammonia in the presence of ATP.

noncompetitive inhibition:  A type of enzyme inhibition not reversed by increasing the substrate concentration.

noncyclic electron flow:  The light-induced flow of electrons from water to NADP+ in oxygen-evolving photosynthesis; it involves both photosystems I and II.

nonessential amino acids:  Amino acids that can be made by humans and other vertebrates from simpler precursors, and are thus not required in the diet.

nonheme iron proteins:  Proteins, usually acting in oxidation–reduction reactions, containing iron but no porphyrin groups.

nonpolar:  Hydrophobic; describing molecules or groups that are poorly soluble in water.

nonsense codon:  A codon that does not specify an amino acid, but signals the termination of a polypeptide chain.

nonsense mutation:  A mutation that results in the premature termination of a polypeptide chain.

nonsense suppressor:  A mutation, usually in the gene for a tRNA, that causes an amino acid to be inserted into a polypeptide in response to a termination codon.

nucleases:  Enzymes that hydrolyze the internucleotide (phosphodiester) linkages of nucleic acids.

nucleic acids:  Biologically occurring polynucleotides in which the nucleotide residues are linked in a specific sequence by phosphodiester bonds; DNA and RNA.

nucleoid:  In bacteria, the nuclear zone that contains the chromosome but has no surrounding membrane.

nucleolus:  A densely staining structure in the nucleus of eukaryotic cells; involved in rRNA synthesis and ribosome formation.

nucleophile:  An electron-rich group with a strong tendency to donate electrons to an electron-deficient nucleus (electrophile); the entering reactant in a bimolecular substitution reaction.

nucleoplasm:  The portion of a cell’s contents enclosed by the nuclear membrane; also called the nuclear matrix.

nucleoside:  A compound consisting of a purine or pyrimidine base covalently linked to a pentose.

nucleoside diphosphate kinase:  An enzyme that catalyzes the transfer of the terminal phosphate of a nucleoside 5′-triphosphate to a nucleoside 5′-diphosphate.

nucleoside diphosphate sugar:  A coenzymelike carrier of a sugar molecule, functioning in the enzymatic synthesis of polysaccharides and sugar derivatives.

nucleoside monophosphate kinase:  An enzyme that catalyzes the transfer of the terminal phosphate of ATP to a nucleoside 5′-monophosphate.

nucleosome:  Structural unit for packaging chromatin; consists of a DNA strand wound around a histone core.

nucleotide:  A nucleoside phosphorylated at one of its pentose hydroxyl groups.

nucleus:  In eukaryotes, a membrane-bounded organelle that contains chromosomes.

oligomer:  A short polymer, usually of amino acids, sugars, or nucleotides; the definition of “short” is somewhat arbitrary, but usually less than 50 subunits.

oligomeric protein:  A multisubunit protein having two or more identical polypeptide chains.

oligonucleotide:  A short polymer of nucleotides (usually less than 50).

oligopeptide:  A few amino acids joined by peptide bonds.

oligosaccharide:  Several monosaccharide groups joined by glycosidic bonds.

oncogene:  A cancer-causing gene; any of several mutant genes that cause cells to exhibit rapid, uncontrolled proliferation. See also proto-oncogene.

open reading frame:  A group of contiguous nonoverlapping nucleotide codons in a DNA or RNA molecule that do not include a termination codon.

open system:  A system that exchanges matter and energy with its surroundings. See also system.

operator:  A region of DNA that interacts with a repressor protein to control the expression of a gene or group of genes.

operon:  A unit of genetic expression consisting of one or more related genes and the operator and promoter sequences that regulate their transcription.

optical activity:  The capacity of a substance to rotate the plane of plane-polarized light.

optimum pH:  The characteristic pH at which an enzyme has maximal catalytic activity.

organelles:  Membrane-bounded structures found in eukaryotic cells; contain enzymes and other components required for specialized cell functions.

origin:  The nucleotide sequence or site in DNA where DNA replication is initiated.

osmosis:  Bulk flow of water through a semipermeable membrane into another aqueous compartment containing solute at a higher concentration.

osmotic pressure:  Pressure generated by the osmotic flow of water through a semipermeable membrane into an aqueous compartment containing solute at a higher concentration.

oxidation:  The loss of electrons from a compound.

oxidation, β:  See β oxidation.

oxidation–reduction reaction:  A reaction in which electrons are transferred from a donor to an acceptor molecule; also called a redox reaction.

oxidative phosphorylation:  The enzymatic phosphorylation of ADP to ATP coupled to electron transfer from a substrate to molecular oxygen.

oxidizing agent (oxidant):  The acceptor of electrons in an oxidation–reduction reaction.

oxygen debt:  The extra oxygen (above the normal resting level) consumed in the recovery period after strenuous physical exertion.

oxygenases:  Enzymes that catalyze reactions in which oxygen is introduced into an acceptor molecule.

palindrome:  A segment of duplex DNA in which the base sequences of the two strands exhibit twofold rotational symmetry about an axis.

paradigm:  In biochemistry, an experimental model or example.

partition coefficient:  A constant that expresses the ratio in which a given solute will be partitioned or distributed between two given immiscible liquids at equilibrium.

pathogenic:  Disease-causing.

pentose:  A simple sugar with a backbone containing five carbon atoms.

pentose phosphate pathway:  A pathway that serves to interconvert hexoses and pentoses and is a source of reducing equivalents and pentoses for biosynthetic processes; present in most organisms. Also called the phosphogluconate pathway.

peptidase:  An enzyme that hydrolyzes a peptide bond.

peptide:  Two or more amino acids covalently joined by peptide bonds.

peptide bond:  A substituted amide linkage between the α-amino group of one amino acid and the α-carboxyl group of another, with the elimination of the elements of water.

peptide mapping:  The characteristic two-dimensional pattern (on paper or gel) formed by the separation of a mixture of peptides resulting from partial hydrolysis of a protein; also known as peptide fingerprinting.

peptidoglycan:  A major component of bacterial cell walls; generally consists of parallel heteropolysaccharides cross-linked by short peptides.

peripheral proteins:  Proteins that are loosely or reversibly bound to a membrane by hydrogen bonds or electrostatic forces; generally water-soluble once released from the membrane.

permeases:  See transporters.

peroxisome:  Membrane-bounded organelle in the cytoplasm of eukaryotic cells; contains peroxide-forming and peroxide-destroying enzymes.

pH:  The negative logarithm of the hydrogen ion concentration of an aqueous solution.

phage:  See bacteriophage.

phenotype:  The observable characteristics of an organism.

phosphodiester linkage:  A chemical grouping that contains two alcohols esterified to one molecule of phosphoric acid, which thus serves as a bridge between them.

phosphogluconate pathway:  An oxidative pathway beginning with glucose-6-phosphate and leading, via 6-phosphogluconate, to pentose phosphates and yielding NADPH. Also called the pentose phosphate pathway.

phospholipid:  A lipid containing one or more phosphate groups.

phosphorolysis:  Cleavage of a compound with phosphate as the attacking group; analogous to hydrolysis.

phosphorylation:  Formation of a phosphate derivative of a biomolecule, usually by enzymatic transfer of a phosphate group from ATP.

phosphorylation potential (ΔGp):  The actual free-energy change of ATP hydrolysis under the nonstandard conditions prevailing within a cell.

photochemical reaction center:  The part of a photosynthetic complex where the energy of an absorbed photon causes charge separation, initiating electron transfer.

photon:  The ultimate unit (a quantum) of light energy.

photophosphorylation:  The enzymatic formation of ATP from ADP coupled to the light-dependent transfer of electrons in photosynthetic cells.

photoreduction:  The light-induced reduction of an electron acceptor in photosynthetic cells.

photorespiration:  Oxygen consumption occurring in illuminated temperate-zone plants, largely due to oxidation of phosphoglycolate.

photosynthesis:  The use of light energy to produce carbohydrates from carbon dioxide and a reducing agent such as water.

photosynthetic phosphorylation:  See photophosphorylation.

photosystem:  In photosynthetic cells, a functional set of light-absorbing pigments and its reaction center.

phototroph:  An organism that can use the energy of light to synthesize its own fuels from simple molecules such as carbon dioxide, oxygen, and water; as distinct from a chemotroph.

pK:  The negative logarithm of an equilibrium constant.

plasma membrane:  The exterior membrane surrounding the cytoplasm of a cell.

plasma proteins:  The proteins present in blood plasma.

plasmalogen:  A phospholipid with an alkenyl ether substituent on the C-1 of glycerol.

plasmid:  An extrachromosomal, independently replicating, small circular DNA molecule; commonly employed in genetic engineering.

plastid:  In plants, a self replicating organelle; may differentiate into a chloroplast.

platelets:  Small, enucleated cells that initiate blood clotting; they arise from cells called megakaryocytes in the bone marrow. Also known as thrombocytes.

pleated sheet:  The side-by-side, hydrogen-bonded arrangement of polypeptide chains in the extended β conformation.

polar:  Hydrophilic, or “water-loving”; describing molecules or groups that are soluble in water.

polarity:  (1) In chemistry, the nonuniform distribution of electrons in a molecule; polar molecules are usually soluble in water. (2) In molecular biology, the distinction between the 5′ and 3′ ends of nucleic acids.

polyclonal antibodies:  A heterogeneous pool of antibodies produced in an animal by a number of different B lymphocytes in response to an antigen. Different antibodies in the pool recognize different parts of the antigen.

polylinker:  A short, often synthetic, fragment of DNA containing recognition sequences for several restriction endonucleases.

polymerase chain reaction (PCR):  A repetitive procedure that results in a geometric amplification of a specific DNA sequence.

polymorphic:  Describing a protein for which amino acid sequence variants exist in a population of organisms, but the variations do not destroy the protein’s function.

polynucleotide:  A covalently linked sequence of nucleotides in which the 3′ hydroxyl of the pentose of one nucleotide residue is joined by a phosphodiester bond to the 5′ hydroxyl of the pentose of the next residue.

polypeptide:  A long chain of amino acids linked by peptide bonds; the molecular weight is generally less than 10,000.

polyribosome:  See polysome.

polysaccharide:  A linear or branched polymer of monosaccharide units linked by glycosidic bonds.

polysome (polyribosome):  A complex of an mRNA molecule and two or more ribosomes.

porphyrin:  Complex nitrogenous compound containing four substituted pyrroles covalently joined into a ring; often complexed with a central metal atom.

positive cooperativity:  A phenomenon of some multisubunit enzymes or proteins in which binding of a ligand or substrate to one subunit facilitates binding to another subunit.

posttranscriptional processing:  The enzymatic processing of the primary RNA transcript, producing functional mRNA, tRNA, and/or rRNA molecules.

posttranslational modification:  Enzymatic processing of a polypeptide chain after translation from its mRNA.

primary structure:  A description of the covalent backbone of a polymer (macromolecule), including the sequence of monomeric subunits and any interchain and intrachain covalent bonds.

primary transcript:  The immediate RNA product of transcription before any posttranscriptional processing reactions.

primase:  An enzyme that catalyzes the formation of RNA oligonucleotides used as primers by DNA polymerases.

primer:  A short oligomer (of sugars or nucleotides, for example) to which an enzyme adds additional monomeric subunits.

probe:  A labeled fragment of nucleic acid containing a nucleotide sequence complementary to a gene or genomic sequence that one wishes to detect in a hybridization experiment.

processivity:  For any enzyme that catalyzes the synthesis of a biological polymer, the property of adding multiple subunits to the polymer without dissociating from the substrate.

prochiral molecule:  A symmetric molecule that can react asymmetrically with an enzyme having an asymmetric active site, generating a chiral product.

projection formulas:  A method for representing molecules to show the configuration of groups around chiral centers; also known as Fischer projection formulas.

prokaryote:  A bacterium; a unicellular organism with a single chromosome, no nuclear envelope, and no membrane-bounded organelles.

promoter:  A DNA sequence at which RNA polymerase may bind, leading to initiation of transcription.

prophage:  A bacteriophage in an inactive state in which the genome is either integrated into the chromosome of the host cell or (sometimes) replicated autonomously.

prostaglandins:  A class of lipid-soluble, hormonelike regulatory molecules derived from arachidonate and other polyunsaturated fatty acids.

prosthetic group:  A metal ion or an organic compound (other than an amino acid) that is covalently bound to a protein and is essential to its activity.

protein:  A macromolecule composed of one or more polypeptide chains, each with a characteristic sequence of amino acids linked by peptide bonds.

protein kinases:  Enzymes that phosphorylate certain amino acid residues in specific proteins.

protein targeting:  The process by which newly synthesized proteins are sorted and transported to their proper locations in the cell.

proteoglycan:  A hybrid macromolecule consisting of a heteropolysaccharide joined to a polypeptide; the polysaccharide is the major component.

proto-oncogene:  A cellular gene, usually encoding a regulatory protein, that can be converted into an oncogene by mutation.

proton acceptor:  An anionic compound capable of accepting a proton from a proton donor; that is, a base.

proton donor:  The donor of a proton in an acid–base reaction; that is, an acid.

proton-motive force:  The electrochemical potential inherent in a transmembrane gradient of H+ concentration; used in oxidative phosphorylation and photophosphorylation to drive ATP synthesis.

protoplasm:  A general term referring to the entire contents of a living cell.

purine:  A nitrogenous heterocyclic base found in nucleotides and nucleic acids; containing fused pyrimidine and imidazole rings.

puromycin:  An antibiotic that inhibits polypeptide synthesis by being incorporated into a growing polypeptide chain, causing its premature termination.

pyranose:  A simple sugar containing the six-membered pyran ring.

pyridine nucleotide:  A nucleotide coenzyme containing the pyridine derivative nicotinamide; NAD or NADP.

pyridoxal phosphate:  A coenzyme containing the vitamin pyridoxine (vitamin B6); functions in reactions involving amino group transfer.

pyrimidine:  A nitrogenous heterocyclic base found in nucleotides and nucleic acids.

pyrimidine dimer:  A covalently joined dimer of two adjacent pyrimidine residues in DNA, induced by absorption of UV light; most commonly derived from two adjacent thymines (a thymine dimer).

pyrophosphatase:  See inorganic pyrophosphatase.

quantum:  The ultimate unit of energy.

quaternary structure:  The three-dimensional structure of a multisubunit protein; particularly the manner in which the subunits fit together.

R group:  (1) Formally, an abbreviation denoting any alkyl group. (2) Occasionally, used in a more general sense to denote virtually any organic substituent (the R groups of amino acids, for example).

racemic mixture (racemate):  An equimolar mixture of the D and L stereoisomers of an optically active compound.

radical:  An atom or group of atoms possessing an unpaired electron; also called a free radical.

radioactive isotope:  An isotopic form of an element with an unstable nucleus that stabilizes itself by emitting ionizing radiation.

radioimmunoassay:  A sensitive and quantitative method for detecting trace amounts of a biomolecule, based on its capacity to displace a radioactive form of the molecule from combination with its specific antibody.

rate-limiting step:  (1) Generally, the step in an enzymatic reaction with the greatest activation energy or the transition state of highest free energy. (2) The slowest step in a metabolic pathway.

reaction intermediate:  Any chemical species in a reaction pathway that has a finite chemical lifetime.

reading frame:  A contiguous and nonoverlapping set of three-nucleotide codons in DNA or RNA.

recombinant DNA:  DNA formed by the joining of genes into new combinations.

redox pair:  An electron donor and its corresponding oxidized form; for example, NADH and NAD+.

redox reaction:  See oxidation-reduction reaction.

reducing agent (reductant):  The electron donor in an oxidation-reduction reaction.

reducing end:  The end of a polysaccharide having a terminal sugar with a free anomeric carbon; the terminal residue can act as a reducing sugar.

reducing equivalent:  A general or neutral term for an electron or an electron equivalent in the form of a hydrogen atom or a hydride ion.

reducing sugar:  A sugar in which the carbonyl (anomeric) carbon is not involved in a glycosidic bond and can therefore undergo oxidation.

reduction:  The gain of electrons by a compound or ion.

regulatory enzyme:  An enzyme having a regulatory function through its capacity to undergo a change in catalytic activity by allosteric mechanisms or by covalent modification.

regulatory gene:  A gene that gives rise to a product involved in the regulation of the expression of another gene; for example, a gene coding for a repressor protein.

regulatory sequence:  A DNA sequence involved in regulating the expression of a gene; for example, a promoter or operator.

regulon:  A group of genes or operons that are coordinately regulated even though some, or all, may be spatially distant within the chromosome or genome.

release factors:  See termination factors.

releasing factors:  Hypothalamic hormones that stimulate release of other hormones by the pituitary gland.

renaturation:  Refolding of an unfolded (denatured) globular protein so as to restore native structure and protein function.

replication:  Synthesis of a daughter duplex DNA molecule identical to the parental duplex DNA.

replisome:  The multiprotein complex that promotes DNA synthesis at the replication fork.

repressible enzyme:  In bacteria, an enzyme whose synthesis is inhibited when its reaction product is readily available to the cell.

repression:  A decrease in the expression of a gene in response to a change in the activity of a regulatory protein.

repressor:  The protein that binds to the regulatory sequence or operator for a gene, blocking its transcription.

residue:  A single unit within a polymer; for example, an amino acid within a polypeptide chain. The term reflects the fact that sugars, nucleotides, and amino acids lose a few atoms (generally the elements of water) when incorporated in their respective polymers.

respiration:  The catabolic process in which electrons are removed from nutrient molecules and passed through a chain of carriers to oxygen.

respiratory chain:  The electron transfer chain; a sequence of electron-carrying proteins that transfer electrons from substrates to molecular oxygen in aerobic cells.

restriction endonucleases:  Site-specific endodeoxyribonucleases causing cleavage of both strands of DNA at points within or near the specific site recognized by the enzyme; important tools in genetic engineering.

restriction fragment:  A segment of double-stranded DNA produced by the action of a restriction endonuclease on a larger DNA.

restriction fragment length polymorphisms (RFLPs):  Variations, among individuals in a population, in the length of certain restriction fragments within which certain genomic sequences occur. These variations result from rare sequence changes that create or destroy restriction sites in the genome.

retrovirus:  An RNA virus containing a reverse transcriptase.

reverse transcriptase:  An RNA-directed DNA polymerase in retroviruses; capable of making DNA complementary to an RNA.

ribonuclease:  A nuclease that catalyzes the hydrolysis of certain internucleotide linkages of RNA.

ribonucleic acid:  See RNA.

ribonucleotide:  A nucleotide containing D-ribose as its pentose component.

ribosomal RNA (rRNA):  A class of RNA molecules serving as components of ribosomes.

ribosome:  A supramolecular complex of rRNAs and proteins, approximately 18 to 22 nm in diameter; the site of protein synthesis.

ribozymes:  Ribonucleic acid molecules with catalytic activities; RNA enzymes.

RNA (ribonucleic acid):  A polyribonucleotide of a specific sequence linked by successive 3′,5′-phosphodiester bonds.

RNA polymerase:  An enzyme that catalyzes the formation of RNA from ribonucleoside 5′-triphosphates, using a strand of DNA or RNA as a template.

RNA splicing:  Removal of introns and joining of exons in a primary transcript.

rRNA:  See ribosomal RNA.

S-adenosylmethionine (adoMet):  An enzymatic cofactor involved in methyl group transfers.

salvage pathway:  Synthesis of a biomolecule, such as a nucleotide, from intermediates in the degradative pathway for the biomolecule; a recycling pathway, as distinct from a de novo pathway.

saponification:  Alkaline hydrolysis of triacylglycerols to yield fatty acids as soaps.

sarcomere:  A functional and structural unit of the muscle contractile system.

satellite DNA:  Highly repeated, nontranslated segments of DNA in eukaryotic chromosomes; most often associated with the centromeric region. Its function is not clear.

saturated fatty acid:  A fatty acid containing a fully saturated alkyl chain.

second law of thermodynamics:  The law stating that in any chemical or physical process, the entropy of the universe tends to increase.

second messenger:  An effector molecule synthesized within a cell in response to an external signal (first messenger) such as a hormone.

secondary metabolism:  Pathways that lead to specialized products not found in every living cell.

secondary structure:  The residue-by-residue conformation of the backbone of a polymer.

sedimentation coefficient:  A physical constant specifying the rate of sedimentation of a particle in a centrifugal field under specified conditions.

Shine-Dalgarno sequence:  A sequence in an mRNA required for binding prokaryotic ribosomes.

shuttle vector:  A recombinant DNA vector that can be replicated in two or more different host species. See also vector.

sickle-cell anemia:  A human disease characterized by defective hemoglobin molecules; caused by a homozygous allele coding for the β chain of hemoglobin.

sickle-cell trait:  A human condition recognized by the sickling of erythrocytes when exposed to low oxygen tension; occurs in individuals heterozygous for the allele responsible for sickle-cell anemia.

signal sequence:  An amino-terminal sequence that signals the cellular fate or destination of a newly synthesized protein.

signal transduction:  The process by which an extracellular signal (chemical, mechanical, or electrical) is amplified and converted to a cellular response.

silent mutation:  A mutation in a gene that causes no detectable change in the biological characteristics of the gene product.

simple diffusion:  The movement of solute molecules across a membrane to a region of lower concentration, unassisted by a protein transporter.

simple protein:  A protein yielding only amino acids on hydrolysis.

site-directed mutagenesis:  A set of methods used to create specific alterations in the sequence of a gene.

site-specific recombination:  A type of genetic recombination that occurs only at specific sequences.

small nuclear RNA (snRNA):  Any of several small RNA molecules in the nucleus; most have a role in the splicing reactions that remove introns from mRNA, tRNA, and rRNA molecules.

somatic cells:  All body cells except the germ-line cells.

SOS response:  In bacteria, a coordinated induction of a variety of genes as a response to high levels of DNA damage.

Southern blot:  A DNA hybridization procedure in which one or more specific DNA fragments are detected in a larger population by means of hybridization to a complementary, labeled nucleic acid probe.

specific activity:  The number of micromoles (μmol) of a substrate transformed by an enzyme preparation per minute per milligram of protein at 25 °C; a measure of enzyme purity.

specific heat:  The amount of energy (in joules or calories) needed to raise the temperature of 1 g of a pure substance by 1 °C.

specific rotation:  The rotation, in degrees, of the plane of plane-polarized light (D-line of sodium) by an optically active compound at 25 °C, with a specified concentration and light path.

specificity:  The ability of an enzyme or receptor to discriminate among competing substrates or ligands.

sphingolipid:  An amphipathic lipid with a sphingosine backbone to which are attached a long-chain fatty acid and a polar alcohol.

splicing:  See gene splicing; RNA splicing.

standard free-energy change (ΔG°):  The free-energy change for a reaction occurring under a set of standard conditions: temperature, 298 K; pressure, 1 atm or 101.3 kPa; and all solutes at 1 M concentration. ΔG°’ denotes the standard free-energy change at pH 7.0.

standard reduction potential (E0′):  The electromotive force exhibited at an electrode by 1 M concentrations of a reducing agent and its oxidized form at 25 °C and pH 7.0; a measure of the relative tendency of the reducing agent to lose electrons.

steady state:  A nonequilibrium state of a system through which matter is flowing and in which all components remain at a constant concentration.

stem cells:  The common, self regenerating cells in bone marrow that give rise to differentiated blood cells such as erythrocytes and lymphocytes.

stereoisomers:  Compounds that have the same composition and the same order of atomic connections, but different molecular arrangements.

sterols:  A class of lipids containing the steroid nucleus.

sticky ends:  Two DNA ends in the same DNA molecule, or in different molecules, with short overhanging single-stranded segments that are complementary to one another, facilitating ligation of the ends; also known as cohesive ends.

stop codons:  See termination codons.

stroma:  The space and aqueous solution enclosed within the inner membrane of a chloroplast, not including the contents within the thylakoid membranes.

structural gene:  A gene coding for a protein or RNA molecule; as distinct from a regulatory gene.

substitution mutation:  A mutation caused by the replacement of one base by another.

substrate:  The specific compound acted upon by an enzyme.

substrate-level phosphorylation:  Phosphorylation of ADP or some other nucleoside 5′-diphosphate coupled to the dehydrogenation of an organic substrate; independent of the electron transfer chain.

suicide inhibitor:  A relatively inert molecule that is transformed by an enzyme, at its active site, into a reactive substance that irreversibly inactivates the enzyme.

suppressor mutation:  A mutation that totally or partially restores a function lost by a primary mutation; located at a site different from the site of the primary mutation.

Svedberg (S):  A unit of measure of the rate at which a particle sediments in a centrifugal field.

symbionts:  Two or more organisms that are mutually interdependent; usually living in physical association.

symport:  Cotransport of solutes across a membrane in the same direction.

synthases:  Enzymes that catalyze condensation reactions in which no nucleoside triphosphate is required as an energy source.

synthetases:  Enzymes that catalyze condensation reactions using ATP or another nucleoside triphosphate as an energy source.

system:  An isolated collection of matter; all other matter in the universe apart from the system is called the surroundings.

telomere:  Specialized nucleic acid structure found at the ends of linear eukaryotic chromosomes.

temperate phage:  A phage whose DNA may be incorporated into the host-cell genome without being expressed; as distinct from a virulent phage, which destroys the host cell.

template:  A macromolecular mold or pattern for the synthesis of an informational macromolecule.

terminal transferase:  An enzyme that catalyzes the addition of nucleotide residues of a single kind to the 3′ end of DNA chains.

termination codons:  UAA, UAG, and UGA; in protein synthesis, signal the termination of a polypeptide chain. Also known as stop codons.

termination factors:  Protein factors of the cytosol required in releasing a completed polypeptide chain from a ribosome; also known as release factors.

termination sequence:  A DNA sequence that appears at the end of a transcriptional unit and signals the end of transcription.

terpenes:  Organic hydrocarbons or hydrocarbon derivatives constructed from recurring isoprene units. They produce some of the scents and tastes of plant products; for example, the scents of geranium leaves and pine needles.

tertiary structure:  The three-dimensional conformation of a polymer in its native folded state.

tetrahydrobiopterin:  The reduced coenzyme form of biopterin.

tetrahydrofolate:  The reduced, active coenzyme form of the vitamin folate.

thiamine pyrophosphate:  The active coenzyme form of vitamin B1; involved in aldehyde transfer reactions.

thioester:  An ester of a carboxylic acid with a thiol or mercaptan.

3′ end:  The end of a nucleic acid that lacks a nucleotide bound at the 3′ position of the terminal residue.

thrombocytes:  See platelets.

thromboxanes:  A class of molecules derived from arachidonate and involved in platelet aggregation during blood clotting.

thylakoid:  Closed cisterna, or disc, formed by the pigment-bearing internal membranes of chloroplasts.

thymine dimer:  See pyrimidine dimer.

tissue culture:  Method by which cells derived from multicellular organisms are grown in liquid media.

titration curve:  A plot of the pH versus the equivalents of base added during titration of an acid.

tocopherols:  Forms of vitamin E.

topoisomerases:  Enzymes that introduce positive or negative supercoils in closed, circular duplex DNA.

topoisomers:  Different forms of a covalently closed, circular DNA molecule that differ only in their linking number.

toxins:  Proteins produced by some organisms and toxic to certain other species.

trace element:  A chemical element required by an organism in only trace amounts.

transaminases:  See aminotransferases.

transamination:  Enzymatic transfer of an amino group from an α-amino acid to an α-keto acid.

transcription:  The enzymatic process whereby the genetic information contained in one strand of DNA is used to specify a complementary sequence of bases in an mRNA chain.

transcriptional control:  The regulation of a protein’s synthesis by regulation of the formation of its mRNA.

transduction:  (1) Generally, the conversion of energy or information from one form to another. (2) The transfer of genetic information from one cell to another by means of a viral vector.

transfer RNA (tRNA):  A class of RNA molecules (Mr 25,000 to 30,000), each of which combines covalently with a specific amino acid as the first step in protein synthesis.

transformation:  Introduction of an exogenous DNA into a cell, causing the cell to acquire a new phenotype.

transgenic:  Describing an organism that has genes from another organism incorporated within its genome as a result of recombinant DNA procedures.

transition state:  An activated form of a molecule in which the molecule has undergone a partial chemical reaction; the highest point on the reaction coordinate.

translation:  The process in which the genetic information present in an mRNA molecule specifies the sequence of amino acids during protein synthesis.

translational control:  The regulation of a protein’s synthesis by regulation of the rate of its translation on the ribosome.

translational repressor:  A repressor that binds to an mRNA, blocking translation.

translocase:  (1) An enzyme that catalyzes membrane transport. (2) An enzyme that causes a movement, such as the movement of a ribosome along an mRNA.

transpiration:  Passage of water from the roots of a plant to the atmosphere via the vascular system and the stomata of the leaves.

transporters:  Proteins that span a membrane and transport specific nutrients, metabolites, ions, or proteins across the membrane; sometimes called permeases.

transposition:  The movement of a gene or set of genes from one site in the genome to another.

transposon (transposable element):  A segment of DNA that can move from one position in the genome to another.

triacylglycerol:  An ester of glycerol with three molecules of fatty acid; also called a triglyceride or neutral fat.

tricarboxylic acid cycle:  See citric acid cycle.

triose:  A simple sugar with a backbone containing three carbon atoms.

tRNA:  See transfer RNA.

tropic hormone (tropin):  A peptide hormone that stimulates a specific target gland to secrete its hormone; for example, thyrotropin produced by the pituitary stimulates secretion of thyroxine by the thyroid.

turnover number:  The number of times an enzyme molecule transforms a substrate molecule per unit time, under conditions giving maximal activity at substrate concentrations that are saturating.

ultraviolet (UV) radiation:  Electromagnetic radiation in the region of 200 to 400 nm.

uncoupling agent:  A substance that uncouples phosphorylation of ADP from electron transfer; for example, 2,4-dinitrophenol.

uniport:  A transport system that carries only one solute, as distinct from cotransport.

unsaturated fatty acid:  A fatty acid containing one or more double bonds.

urea cycle:  A metabolic pathway in vertebrates, for the synthesis of urea from amino groups and carbon dioxide; occurs in the liver.

ureotelic:  Excreting excess nitrogen in the form of urea.

uricotelic:  Excreting excess nitrogen in the form of urate (uric acid).

Vmax:  The maximum velocity of an enzymatic reaction when the binding site is saturated with substrate.

vector:  A DNA molecule known to replicate autonomously in a host cell, to which a segment of DNA may be spliced to allow its replication; for example, a plasmid or a temperate-phage DNA.

viral vector:  A viral DNA altered so that it can act as a vector for recombinant DNA.

virion:  A virus particle.

virus:  A self replicating, infectious, nucleic acid–protein complex that requires an intact host cell for its replication; its genome is either DNA or RNA.

vitamin:  An organic substance required in small quantities in the diet of some species; generally functions as a component of a coenzyme.

wild type:  The normal (unmutated) phenotype.

wobble:  The relatively loose base pairing between the base at the 3′ end of a codon and the complementary base at the 5′ end of the anticodon.

x-ray crystallography:  The analysis of x-ray diffraction patterns of a crystalline compound, used to determine the molecule’s three-dimensional structure.

zinc finger:  A specialized protein motif involved in DNA recognition by some DNA-binding proteins; characterized by a single atom of zinc coordinated to four Lys residues or to two His and two Lys residues.

zwitterion:  A dipolar ion, with spatially separated positive and negative charges.

zymogen:  An inactive precursor of an enzyme; for example, pepsinogen, the precursor of pepsin.
Chapter H
Hall of Fame
Hodgkin, Dorothy Crowfoot, 495
Lavoisier, Antoine, 56, 364
Schrödinger, Erwin, 4
Szent-Györgyi, Albert, 461
Chapter N

Page numbers in boldface type indicate where a structural formula is given.

abiotic [synthesis] (3)
absolute configuration (7)
absolute temperature (4)
acetic acid (40)
       pKa, 96
       titration curve, 95–96
acetic acid–acetate pair, 94
acetylcholine (19)
       molecular weight, 31
N-acetylgalactosamine (1)
N-acetylmuramic acid (5)
N-acetylneuraminic acid (2), 305
       definition, 94
adaptations, biochemical (1)
adenine (7), 71, 325
       in DNA, 325
       in RNA, 325
adenosine 5′-diphosphate (ADP) (20), 67
       in ATP cycle, 374–375, 378–379, 381
adenosine 2′-monophosphate, 327
adenosine 3′-monophosphate, 327
adenosine 5′-monophosphate (AMP), 326, (see also adenylate)
       from ATP, 484
       in ATP cycle, 378–379, 381
alanine (21), 70, 115
alcohol (20)
alkalosis (1)
D-allose, 301
allosteric site[s] (3)
α-amino [acids] (28), 70
α/β barrel (2)
α carbon (10)
α helix (47)
       hydrogen bonding, 167
       in proteins, 161, 163, 166–169
       right- vs. left-handed, 167
       stability and amino acid sequence, 168–169
       stereoisomers in, 167
D-altrose, 301
Amanita phalloides (1)
amino-acid analyzer (1)
antigen–antibody complex (1)
antigen[s] (26)
apoenzyme (1)
apurinic acid (1), 344
D-arabinose, 301
L-arabinose, 302
arachidonic acid (10), (see also arachidonate)
archaebacteria (4)
       membranes of, 27
L-ascorbic acid (5), 438, (see also vitamin C)
       deficiency, 439
       synthesis, 437–439
Bacillus brevis (3)
Bacillus subtilis (1)
βαβ loop (2)
β barrel (1)
β bend (3)
β conformation, in proteins (18)
β pleated sheet (5)
β turn (7)
bicarbonate (23)
       in blood plasma, 100–101
       buffer system, 100–101
       chloride–bicarbonate exchanger, 278, 281, 286
biosynthesis (16), see anabolism
blood (124), (see also blood plasma)
buffer[s] (95), 96
cancer (3)
carbohydrates (42), 298–320, (see also disaccharides; monosaccharides; oligosaccharides; polysaccharides)
       abundance, 298
       analysis of, 318
       complex, 298
       empirical formulas, 298
       in glycolipids, 304–305, 315–316, 318
       in glycoproteins, 302, 304–305, 315–318
       size classes, 298
       synthesis, 305, (see also carbohydrate biosynthesis)
carbon monoxide, 26
γ-carboxyglutamate (3), 118
Chlamydomonas (4)
chloride–bicarbonate exchanger (4)
cholesterol (17), 254
choline, 71
chondroitin sulfate (2), 313
chromatin (15)
       fibers, size of, 37
chromosomes (25)
       proteins in, 37
chymotrypsinogen (5)
       amino acid residues, 136–137
cilia (23)
       ATP-driven motion, 45
       microtubule construction, 45
cisternae, 32–34, 39
colchicine (3)
collagen (50)
condensation reactions (6)
configuration, definition, 63
configurational isomers (1)
conformation (172)
       definition, 64
       eclipsed, 64
       entropy of, 162
       monosaccharides, 304
       of proteins, 17–18, 154, 160–161
       staggered, 64
cytosine (1), 71, 325
dabsyl chloride (6), 124
defense protein (1)
dehydrogenase[s] (7)
denaturation (9)
deoxyadenosine 5′-monophosphate, 326
deoxyadenosylcobalamin, 200, (see also coenzyme B12)
deoxyadenylate (1), 326
deoxycytidine 5′-monophosphate (dCMP), 326
deoxycytidylate (1), 326
2-deoxy-D-ribose (2), 299
2-deoxy-α-D-ribose, 71
2′-deoxy-D-ribose (in DNA) (1)
deoxyguanylate (1), 326
deoxyhemoglobin, 188
Dictyostelium (1)
dideoxy sequencing, 347–349
digestion (6)
dihydroxyacetone (2), 299, 301
1,25-dihydroxycholecalciferol (2), 260
dimethylsulfate, as mutagen (1), 346
diphtheria toxin (2)
disaccharides, 298
       O-glycosidic bond (3), 306
       N-glycosidic bond[s] (3)
       nomenclature rules, 306–307
       reducing end, 306
dissociation constant[s] (10)
       for ES complex (KS), 216, 224, 228–229
       for weak acids, 94
disulfide bond[s] (cross-links) (56)
       cleavage, 150
       in cystine, 116, 150
double lactim tautomer, 330
double-reciprocal plot (8)
dynamic steady state (6)
dynein (11)
E. coli (38), see Escherichia coli
earth, age of, 76
eclipsed conformation (1)
egg, lysozyme in, 180
egg albumin, denaturation, 180
Ehlers–Danlos syndrome (1)
EI (enzyme–inhibitor) complex, 220
elastase (1)
elastin (12)
entropy (S) (31)
enzymes, 198–346, (see also catalysis)
       active site (56)
       binding energy [in catalysis] (32)
       as catalysts, 10–12, 198, 202–211
       catalytic functional groups, 205
       chiral compounds, 64, 114
       classification and naming, 200–201
       coenzymes, see coenzyme
       cofactor[s] (60)
       conformational change, 182, 208
       denaturation, 199
D-erythrose, 301
Escherichia coli (6)
Euglena (2)
evolutionary maps (1)
exergonic [reactions] (37)
exothermic [reaction] (3)
extracellular matrix (12)
extrinsic [membrane proteins] (1), (see also peripheral membrane proteins)
Faraday constant ( ℱ ) (3)
fat cells (2), (see also adipocytes)
fats (21), (see also triacylglycerols)
       in adipose tissue, 480
       in diet, 497
ferritin (1)
filamin (3)
“flickering clusters” (1)
flip-flop diffusion (5)
fluid mosaic [model] (7)
fluorescamine reaction, 124
1-fluoro-2,4-dinitrobenzene (FDNB) (8), 124, 149
fodrin (2)
fossil[s] (9)
free energy (G) (97)
       thermodynamic definition, 72
       use by cells, 71, 368
free-energy change (ΔG) (113), (see also standard free-energy change)
       actual, 370–372
       entropy change and, 366–367
       standard, 91, 202, 204, 287, 368
       in transport processes, 284, 287, 383
ΔG (29), see free-energy change
gene expression (2)
genetic counseling (1)
Gentian violet (1)
geometric isomers (1)
Gibbs free energy (2), see free energy
glial cells (1)
glial fibrillary acidic protein (1)
gluconate, 305, 306
gluconic acid (1)
α-D-glucopyranose (3), 302, 304
β-D-glucopyranose (3), 302
glucosamine (4), 305
D-glucose, 67, 299, 301, 307
glutamic acid (1), 70, (see also glutamate)
glutamine (6), 70, 115
glycans, 308, (see also polysaccharides)
glyceraldehyde (8), 299
       optical activity, 300
       stereoisomers, 113
glycine (66), 70, 115
       as buffer, 119
       isoelectric point, 120
       pKa, 119
       titration curve, 118–119
glycoconjugates (4), 315
glycogen (28)
       composition, 310
       conformation, 311
       as energy source, 244
growth factor[s] (1)
D-gulose, 301
H (enthalpy) (5)
ΔH (enthalpy change) (17)
HA (fusion protein), 282
Halobacterium (1), ~ halobium (2)
haploid [cells] (2)
Haworth perspective [formulas] (6)
heat [energy] (7)
heat-shock proteins, 184
hemiacetal[s] (12)
hemiketal[s] (4)
hemoglobin (88)
       conformational changes, 182, 188–189
       in evolution, 186
human rhinovirus (1)
hyaluronidase (1)
hybridization techniques, 343
hydration (11)
hydrogen, in living matter, 57
hydrogen bonds (91), (see also weak interactions)
       in α helix, 167
       bond energy, 82, 88
       in DNA, 331, 334–335, 821
5-hydroxylysine (2), 118
4-hydroxyproline (2), 118
D-idose, 301
inflammation (4)
information (102), (see also genetic information)
intrinsic membrane proteins, 277–279, (see also integral membrane proteins)
invariant residue[s] (4)
inverted repetitions, in DNA (1)
in vitro studies, limitation of, 47–48
ion-exchange chromatography (9)
       of amino acids, 122
       of proteins, 138
ion gradients (9)
ionic interactions (21), (see also weak interactions)
ionization (43)
Isoetes muricata (1)
isoleucine (23), 12, 70, 115
isoprene (10), 256
kinesin (6)
lactam tautomer, 330
lactim tautomer, 330
lactones (1)
lactose (17), 307
Lambert–Beer law (3)
laminin (1)
lanolin (1)
lauric acid (1)
leather (1)
leucine (4), 70, 115
levorotation (L), 113
ligand (17)
Lineweaver–Burk equation (2)
linoleoyl-CoA (1), 492
lysine (8), 70, 115
lysinonorleucine (3)
lysophospholipase[s] (2)
lysophospholipid[s] (2)
lysosomes (26)
       function, 34
       lysosomal membrane, 34
       pH, 34
       size, 34
maleic acid (3), 63
maltose (9), 307
mannosamine (1), 305
metabolism (48)
       aerobic (3)
       for balance and economy, 13
metabolites (11)
metal-ion catalysis, 210
metalloproteins (1)
methane (5)
       diffusion across membranes, 284
       in early atmosphere, 26, 74
methionine (4), 70, 115
N6-methyladenine (3), 327
methylation analysis (of carbohydrates) (1), 318
5-methylcytosine (4), 327
N2-methylguanine, 327
N-methyllysine (1), 118
mycoplasma (5)
Mycoplasma pneumoniae (1)
myoglobin (57)
       amino acid residues, 178
       amino acid sequence, 176–177, 181
       in diving mammals, 176–177, 181
       in evolution, 186
       heme group, 176, 178–179
       and hemoglobin, 186
       hydrophobic core, 178
       oxygen-saturation curve, 188
       oxygen storage, 176, 189
native protein[s] (2)
Neisseria gonorrhoeae (1)
Neurospora (1)
Nitella (4)
nuclear magnetic resonance (NMR) spectroscopy (3)
nucleoid (11)
nucleolus (6)
       in mitosis, 38
       RNA content, 37
nucleophile[s] (10)
nucleoplasm (4)
nucleoside (22)
       definition, 325
       nomenclature, 325, 350–351
nucleoside 5′-monophosphate (NMP) (3), 351
nucleoside 5′-triphosphate (NTP) (1), 351
nucleosome[s] (7)
oleic acid (5), 71, 242, (see also oleate)
Ophioglossum (1)
palmitic acid (8), 71, (see also palmitate)
Paramecium (5)
parasite (6)
       viral, 50
paratartaric acid (1)
partial pressure (8)
       of oxygen (pO2), 188–189
passive transport (9)
pentoses (11)
       aldo and keto forms, 299
       in nucleic acids, 299, 326
pepsin (9)
       crystallization of, 199
       molecular weight, 509
phospholipase A (3)
phospholipase D, 264, 280
phospholipases (9)
phosphonate [compounds] (1)
phosphorylation potential (2)
photoautotrophs (2)
photoheterotrophs (1)
phototroph[s] (2)
phylogenetic tree (1)
       homologous proteins and, 155
phylogeny, 17, 155
plasmids (2)
plastids (2)
platelet-activating factor (5)
positive cooperativity (2)
potassium (K)
       cotransport with sodium, 288–289
potential energy (11)
ppGpp (1), see guanosine 3′-diphosphate,5′-diphosphate
pre-steady state [kinetics] (7)
primary structure (8)
       of nucleic acids, 331
       of proteins, 134, 146, 153, 161
priming reactions, 406, 407
“primordial soup” (4)
procollagen (1)
proline (17), 70, 115
prosthetic group[s] (10)
proteoglycans (18)
       composition, 313
       core proteins, 313–314
protist[s] (15)
protofibril (2)
protofilaments (2)
protomers (1)
proton acceptor (22)
proton donor (27)
proton pump (7)
pseudouracil (2), 327
P-type ATPase (4)
purine, 325
purine bases, 325
pyranoses (3)
quaternary structure (15)
racemic acid (1)
racemic mixture (3)
Ramachandran plot (5)
rancid[ity] (3)
rate constant (k) (18)
rate law (3)
reaction coordinate diagram (8)
reaction intermediate[s] (5)
reaction rates, 11, 202–204, 352, 372
receptor ligand (1)
recognition sites, 252
red blood cell, 25, (see also erythrocyte)
reducing end, of disaccharide, 306
retinol (3), 259, (see also vitamin A)
rhamnose (1), 305
rhodamine (1)
Rhodopseudomonas viridis (1)
rhodopsin (7)
ribonucleic acid (RNA) (245), (see also messenger RNA, ribosomal RNA, transfer RNA)
       antiparallel strands, 339
       base composition, 325
       base pairing, 339
       base-stacking interactions, 339–340
       in chloroplasts, 39
       classes of, 324
       denaturation, 342–343
D-ribose (4), 299, 301
α-D-ribose, 71
D-ribulose (2), 301
rotation (50)
       around bonds, 64, 165, 311
       of plane-polarized light (11)
rough endoplasmic reticulum (RER) (10)
rumen, 311
ΔS (entropy change) (14)
[S]0.5 (1)
S. typhimurium, see Salmonella
Saccharomyces (2), ~ cerevisiae (1)
saddle [shape] (4)
Salmonella, 318
selenocysteine (2), 118
self-replica[tion] (14), 76
slime molds (2)
soap[s] (9)
sodium dodecylsulfate (SDS) (15), 141
solvation shell (1)
space-filling model[s] (14)
specific acid or base catalysis (1)
specific activity (enzymes), 140
specificity (enzymes) (27)
spectrin (3)
spermaceti oil (8)
spermaceti organ (2)
Staphylococcus aureus (1)
starch (11)
stearic acid (2), 242, (see also stearate)
steroid[s] (18)
system, definition of, 8
D-talose, 301
tartaric acid (4)
termites (1)
ternary complex (6)
tertiary structure (37)
       of nucleic acids, 331
       of proteins, 161, 175–185
Tetrahymena (1)
tetroses (4)
thermodynamics (26), 85, 183, (see also enthalpy; entropy; free energy)
       bioenergetics and, 365–373
D-threose, 301
thyrotropin-releasing factor [hormone] (2), 127
transition state (63)
transition-state stabilization (1)
transition-state theory (1)
transpiration (2)
transport ATPases (3)
       ATP synthase, 290
       F-type, 290
       inhibition, 289
       Na+K+ ATPase, 288–289, 292, 383
       P-type, 289–290
trehalose (2), 307
tyrosine (15), 70, 115
ubiquinone (coenzyme Q, UQ) (5), 262
ubiquitin (2)
uncompetitive inhibitor (4)
unicellular organism[s] (6)
uniport systems (1)
universe, defined, 8
uracil (5), 71, 325
urease (8)
       crystallization of, 160, 199
       molecular weight, 160
uridine diphosphate (UDP), 423
       carrier of hexose groups, 423
V0 (initial velocity) (54)
valine (5), 70, 115
valinomycin (5)
vitalism (1)
vitamin B1, 200
water (380), 81–104
       abundance in cells, 68, 81
       dielectric constant, 84
       diffusion across membranes, 284
       geometry, 82
       as heat buffer, 103
       hydrogen bonding, 81–83, 88, 162–163
       internal cohesion, 82–83, 103
       ionization, 81, 90–92
       ion product, 91–92
       “metabolic water”, 102, 359, 536
wax[es] (19)
wild-type cells (2)
D-xylose, 301
yeast (7)

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* 586,662

Chapter R

I p:        __________________________________________________ ?
II p:        __________________________________________________ ?
III p:        __________________________________________________ ?
j: Figure 1       ____________________________________________ ?
m: Figure 2       ____________________________________________ ?
m: Figure 3       ____________________________________________ ?
n: Figure 4       ____________________________________________ ?
13 r: Antoine Lavoisier        ____________________________________ ?
13.1.1 –: “Now, in the second law of thermodynamics . . .” _________ ?
–: Table 13–1   ____________________________________________ ?
Box 13–1  k: The oxidation of glucose      _______________________________ ?
k: Letters fallen into a completely random, chaotic pattern    ______ ?
13.1.3 –: Table 13–2   ____________________________________________ ?
–: Table 13–3   ____________________________________________ ?
n: !Table 13–4  ____________________________________________ 76,334
13.2.1 m: Figure 13–1       _________________________________________ ?
m: Figure 13–2       _________________________________________ ?
n: !Table 13–5  ____________________________________________ ?
13.2.2 m: Figure 13–3       _________________________________________ ?
m: Figure 13–4       _________________________________________ ?
m: Figure 13–6       _________________________________________ ?
m: Figure 13–7       _________________________________________ ?
m: Figure 13–5       _________________________________________ ?
n: !Table 13–6  ____________________________________________ ?
13.2.3 m: Figure 13–8       _________________________________________ ?
m: Figure 13–9       _________________________________________ ?
m: Figure 13–10       ________________________________________ ?
13.2.4 m: Figure 13–11       ________________________________________ ?
Box 13–3  m: Figure 1       ____________________________________________ ?
n: Figure 2       ____________________________________________ 32,334
13.2.6 m: Figure 13–12       ________________________________________ ?
13.3.1 m: Figure 13–13       ________________________________________ ?
13.3.2 k: The oxidation of a reducing sugar by cupric ion        ___________ ?
k: The above reaction expressed as two half reactions   __________ ?
13.3.3 m: Figure 13–14       ________________________________________ ?
13.3.4 m: Figure 13–15       ________________________________________ ?
–: Table 13–7   ____________________________________________ ?
k: Eqn (13–6)   ____________________________________________ ?
k: Eqn (13–7)   ____________________________________________ ?
13.3.5 k: Reduction protential calculation   ___________________________ 16,192
13.3.8 n: Figure 13–16       ________________________________________ 71,469