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Alberts B, Johnson A, Lewis J, et al. Molecular Biology of the Cell. 4th edition. New York: Garland Science; 2002.
We have seen that each type of protein consists of a precise sequence of amino acids that allows it to fold up into a particular three-dimensional shape, or conformation. But proteins are not rigid lumps of material. They can have precisely engineered moving parts whose mechanical actions are coupled to chemical events. It is this coupling of chemistry and movement that gives proteins the extraordinary capabilities that underlie the dynamic processes in living cells.
In this section, we explain how proteins bind to other selected molecules and how their activity depends on such binding. We show that the ability to bind to other molecules enables proteins to act as catalysts, signal receptors, switches, motors, or tiny pumps. The examples we discuss in this chapter by no means exhaust the vast functional repertoire of proteins. However, the specialized functions of many of the proteins you will encounter elsewhere in this book are based on similar principles.
The biological properties of a protein molecule depend on its physical interaction with other molecules. Thus, antibodies attach to viruses or bacteria to mark them for destruction, the enzyme hexokinase binds glucose and ATP so as to catalyze a reaction between them, actin molecules bind to each other to assemble into actin filaments, and so on. Indeed, all proteins stick, or bind, to other molecules. In some cases, this binding is very tight; in others, it is weak and short-lived. But the binding always shows great specificity, in the sense that each protein molecule can usually bind just one or a few molecules out of the many thousands of different types it encounters. The substance that is bound by the protein—no matter whether it is an ion, a small molecule, or a macromolecule— is referred to as a ligand for that protein (from the Latin word ligare, meaning “to bind”).
The ability of a protein to bind selectively and with high affinity to a ligand depends on the formation of a set of weak, noncovalent bonds—hydrogen bonds, ionic bonds, and van der Waals attractions—plus favorable hydrophobic interactions (see Panel 2-3, pp. 114–115). Because each individual bond is weak, an effective binding interaction requires that many weak bonds be formed simultaneously. This is possible only if the surface contours of the ligand molecule fit very closely to the protein, matching it like a hand in a glove (Figure 3-37).
The selective binding of a protein to another molecule. Many weak bonds are needed to enable a protein to bind tightly to a second molecule, which is called a ligand for the protein. A ligand must therefore fit precisely into a protein’s binding (more...)
The region of a protein that associates with a ligand, known as the ligand’s binding site, usually consists of a cavity in the protein surface formed by a particular arrangement of amino acids. These amino acids can belong to different portions of the polypeptide chain that are brought together when the protein folds (Figure 3-38). Separate regions of the protein surface generally provide binding sites for different ligands, allowing the protein’s activity to be regulated, as we shall see later. And other parts of the protein can serve as a handle to place the protein in a particular location in the cell—an example is the SH2 domain discussed previously, which is often used to move a protein containing it to sites in the plasma membrane in response to particular signals.
The binding site of a protein. (A) The folding of the polypeptide chain typically creates a crevice or cavity on the protein surface. This crevice contains a set of amino acid side chains disposed in such a way that they can make noncovalent bonds only (more...)
Although the atoms buried in the interior of the protein have no direct contact with the ligand, they provide an essential scaffold that gives the surface its contours and chemical properties. Even small changes to the amino acids in the interior of a protein molecule can change its three-dimensional shape enough to destroy a binding site on the surface.
Proteins have impressive chemical capabilities because the neighboring chemical groups on their surface often interact in ways that enhance the chemical reactivity of amino acid side chains. These interactions fall into two main categories.
First, neighboring parts of the polypeptide chain may interact in a way that restricts the access of water molecules to a ligand binding site. Because water molecules tend to form hydrogen bonds, they can compete with ligands for sites on the protein surface. The tightness of hydrogen bonds (and ionic interactions) between proteins and their ligands is therefore greatly increased if water molecules are excluded. Initially, it is hard to imagine a mechanism that would exclude a molecule as small as water from a protein surface without affecting the access of the ligand itself. Because of the strong tendency of water molecules to form water–water hydrogen bonds, however, water molecules exist in a large hydrogen-bonded network (see Panel 2-2, pp. 112–113). In effect, a ligand binding site can be kept dry because it is energetically unfavorable for individual water molecules to break away from this network, as they must do to reach into a crevice on a protein’s surface.
Second, the clustering of neighboring polar amino acid side chains can alter their reactivity. If a number of negatively charged side chains are forced together against their mutual repulsion by the way the protein folds, for example, the affinity of the site for a positively charged ion is greatly increased. In addition, when amino acid side chains interact with one another through hydrogen bonds, normally unreactive side groups (such as the –CH2OH on the serine shown in Figure 3-39) can become reactive, enabling them to enter into reactions that make or break selected covalent bonds.
An unusually reactive amino acid at the active site of an enzyme. This example is the “catalytic triad” found in chymotrypsin, elastase, and other serine proteases (see Figure 3-14). The aspartic acid side chain (Asp 102) induces the histidine (more...)
The surface of each protein molecule therefore has a unique chemical reactivity that depends not only on which amino acid side chains are exposed, but also on their exact orientation relative to one another. For this reason, even two slightly different conformations of the same protein molecule may differ greatly in their chemistry.
As we have described previously, many of the domains in proteins can be grouped into families that show clear evidence of their evolution from a common ancestor, and genome sequences reveal large numbers of proteins that contain one or more common domains. The three-dimensional structures of the members of the same domain family are remarkably similar. For example, even when the amino acid sequence identity falls to 25%, the backbone atoms in a domain have been found to follow a common protein fold within 0.2 nanometers (2 Å).
These facts allow a method called “evolutionary tracing” to be used to identify those sites in a protein domain that are the most crucial to the domain’s function. For this purpose, those amino acids that are unchanged, or nearly unchanged, in all of the known protein family members are mapped onto a structural model of the three-dimensional structure of one family member. When this is done, the most invariant positions often form one or more clusters on the protein surface, as illustrated in Figure 3-40A for the SH2 domain described previously (see Panel 3-2, pp. 138–139). These clusters generally correspond to ligand binding sites.
The evolutionary trace method applied to the SH2 domain. (A) Front and back views of a space-filling model of the SH2 domain, with evolutionarily conserved amino acids on the protein surface colored yellow, and those more toward the protein interior colored (more...)
The SH2 domain is a module that functions in protein–protein interactions. It binds the protein containing it to a second protein that contains a phosphorylated tyrosine side chain in a specific amino acid sequence context, as shown in Figure 3-40B. The amino acids located at the binding site for the phosphorylated polypeptide have been the slowest to change during the long evolutionary process that produced the large SH2 family of peptide recognition domains. Because mutation is a random process, this result is attributed to the preferential elimination during evolution of all organisms whose SH2 domains became altered in a way that inactivated the SH2-binding site, thereby destroying the function of the SH2 domain.
In this era of extensive genome sequencing, many new protein families have been discovered whose functions are unknown. By identifying the critical binding sites on a three-dimensional structure determined for one family member, the above method of evolutionary tracing is being used to help determine the functions of such proteins.
Proteins can bind to other proteins in at least three ways. In many cases, a portion of the surface of one protein contacts an extended loop of polypeptide chain (a “string”) on a second protein (Figure 3-41A). Such a surface–string interaction, for example, allows the SH2 domain to recognize a phosphorylated polypeptide as a loop on a second protein, as just described, and it also enables a protein kinase to recognize the proteins that it will phosphorylate (see below).
Three ways in which two proteins can bind to each other. Only the interacting parts of the two proteins are shown. (A) A rigid surface on one protein can bind to an extended loop of polypeptide chain (a “string”) on a second protein. (B) (more...)
A second type of protein–protein interface is formed when two α helices, one from each protein, pair together to form a coiled-coil (Figure 3-41B). This type of protein interface is found in several families of gene regulatory proteins, as discussed in Chapter 7.
The most common way for proteins to interact, however, is by the precise matching of one rigid surface with that of another (Figure 3-41C). Such interactions can be very tight, since a large number of weak bonds can form between two surfaces that match well. For the same reason, such surface–surface interactions can be extremely specific, enabling a protein to select just one partner from the many thousands of different proteins found in a cell.
All proteins must bind to particular ligands to carry out their various functions. This capacity for tight selective binding is displayed to an extraordinary degree by the antibody family, as discussed in detail in Chapter 24.
Antibodies, or immunoglobulins, are proteins produced by the immune system in response to foreign molecules, such as those on the surface of an invading microorganism. Each antibody binds to a particular target molecule extremely tightly, thereby either inactivating the target directly or marking it for destruction. An antibody recognizes its target (called an antigen) with remarkable specificity. Because there are potentially billions of different antigens we might encounter, we have to be able to produce billions of different antibodies.
Antibodies are Y-shaped molecules with two identical binding sites that are complementary to a small portion of the surface of the antigen molecule. A detailed examination of the antigen-binding sites of antibodies reveals that they are formed from several loops of polypeptide chain that protrude from the ends of a pair of closely juxtaposed protein domains (Figure 3-42). The enormous diversity of antigen-binding sites possessed by different antibodies is generated by changing only the length and amino acid sequence of these loops, without altering the basic protein structure.
An antibody molecule. (A) A typical antibody molecule is Y-shaped and has two identical binding sites for its antigen, one on each arm of the Y. The protein is composed of four polypeptide chains (two identical heavy chains and two identical and smaller (more...)
Loops of this kind are ideal for grasping other molecules. They allow a large number of chemical groups to surround a ligand so that the protein can link to it with many weak bonds. For this reason, loops are often used to form the ligand-binding sites in proteins.
Molecules in the cell encounter each other very frequently because of their continual random thermal movements. When colliding molecules have poorly matching surfaces, few noncovalent bonds form, and the two molecules dissociate as rapidly as they come together. At the other extreme, when many noncovalent bonds form, the association can persist for a very long time (Figure 3-43). Strong interactions occur in cells whenever a biological function requires that molecules remain associated for a long time—for example, when a group of RNA and protein molecules come together to make a subcellular structure such as a ribosome.
How noncovalent bonds mediate interactions between macromolecules.
The strength with which any two molecules bind to each other can be measured directly. As an example, imagine a situation in which a population of identical antibody molecules suddenly encounters a population of ligands diffusing in the fluid surrounding them. At frequent intervals, one of the ligand molecules will bump into the binding site of an antibody and form an antibody–ligand complex. The population of antibody–ligand complexes will therefore increase, but not without limit: over time, a second process, in which individual complexes break apart because of thermally induced motion, will become increasingly important. Eventually, any population of antibody molecules and ligands will reach a steady state, or equilibrium, in which the number of binding (association) events per second is precisely equal to the number of “unbinding” (dissociation) events (see Figure 2-52).
From the concentrations of the ligand, antibody, and antibody–ligand complex at equilibrium, one can calculate a convenient measure—termed the equilibrium constant (K)—of the strength of binding (Figure 3-44A). The equilibrium constant is greater the greater the binding strength, and it is a direct measure of the free-energy difference between the bound and free states (Figure 3-44B). Even a change of a few noncovalent bonds can have a striking effect on a binding interaction, as shown by the example in Figure 3-44C.
Relating binding energies to the equilibrium constant. (A) The equilibrium between molecules A and B and the complex AB is maintained by a balance between the two opposing reactions shown in panels 1 and 2. Molecules A and B must collide if they are to (more...)
We have used the case of an antibody binding to its ligand to illustrate the effect of binding strength on the equilibrium state, but the same principles apply to any molecule and its ligand. Many proteins are enzymes, which, as we now discuss, first bind to their ligands and then catalyze the breakage or formation of covalent bonds in these molecules.
Many proteins can perform their function simply by binding to another molecule. An actin molecule, for example, need only associate with other actin molecules to form a filament. There are other proteins, however, for which ligand binding is only a necessary first step in their function. This is the case for the large and very important class of proteins called enzymes. As described in Chapter 2, enzymes are remarkable molecules that determine all the chemical transformations that make and break covalent bonds in cells. They bind to one or more ligands, called substrates, and convert them into one or more chemically modified products, doing this over and over again with amazing rapidity. Enzymes speed up reactions, often by a factor of a million or more, without themselves being changed—that is, they act as catalysts that permit cells to make or break covalent bonds in a controlled way. It is the catalysis of organized sets of chemical reactions by enzymes that creates and maintains the cell, making life possible.
Enzymes can be grouped into functional classes that perform similar chemical reactions (Table 3-1). Each type of enzyme within such a class is highly specific, catalyzing only a single type of reaction. Thus, hexokinase adds a phosphate group to d-glucose but ignores its optical isomer l-glucose; the blood-clotting enzyme thrombin cuts one type of blood protein between a particular arginine and its adjacent glycine and nowhere else, and so on. As discussed in detail in Chapter 2, enzymes work in teams, with the product of one enzyme becoming the substrate for the next. The result is an elaborate network of metabolic pathways that provides the cell with energy and generates the many large and small molecules that the cell needs (see Figure 2-35).
Some Common Types of Enzymes.
For a protein that catalyzes a chemical reaction (an enzyme), the binding of each substrate molecule to the protein is an essential prelude. In the simplest case, if we denote the enzyme by E, the substrate by S, and the product by P, the basic reaction path is E + S → ES → EP → E + P. From this reaction path, we see that there is a limit to the amount of substrate that a single enzyme molecule can process in a given time. If the concentration of substrate is increased, the rate at which product is formed also increases, up to a maximum value (Figure 3-45). At that point the enzyme molecule is saturated with substrate, and the rate of reaction (Vmax) depends only on how rapidly the enzyme can process the substrate molecule. This maximum rate divided by the enzyme concentration is called the turnover number. The turnover number is often about 1000 substrate molecules processed per second per enzyme molecule, although turnover numbers between 1 and 10,000 are known.
Enzyme kinetics. The rate of an enzyme reaction (V) increases as the substrate concentration increases until a maximum value (Vmax) is reached. At this point all substrate-binding sites on the enzyme molecules are fully occupied, and the rate of reaction (more...)
The other kinetic parameter frequently used to characterize an enzyme is its Km, the concentration of substrate that allows the reaction to proceed at one-half its maximum rate (0.5 Vmax) (see Figure 3-45). A low Km value means that the enzyme reaches its maximum catalytic rate at a low concentration of substrate and generally indicates that the enzyme binds to its substrate very tightly, whereas a high Km value corresponds to weak binding. The methods used to characterize enzymes in this way are explained in Panel 3-3 (pp. 164–165).
Some of the Methods Used to Study Enzymes.
Extremely high rates of chemical reaction are achieved by enzymes—far higher than for any synthetic catalysts. This efficiency is attributable to several factors. The enzyme serves, first, to increase the local concentration of substrate molecules at the catalytic site and to hold all the appropriate atoms in the correct orientation for the reaction that is to follow. More importantly, however, some of the binding energy contributes directly to the catalysis. Substrate molecules must pass through a series of intermediate states of altered geometry and electron distribution before they form the ultimate products of the reaction. The free energy required to attain the most unstable transition state is called the activation energy for the reaction, and it is the major determinant of the reaction rate. Enzymes have a much higher affinity for the transition state of the substrate than they have for the stable form. Because this tight binding greatly lowers the energies of the transition state, the enzyme greatly accelerates a particular reaction by lowering the activation energy that is required (Figure 3-46).
Enzymatic acceleration of chemical reactions by decreasing the activation energy. Often both the uncatalyzed reaction (A) and the enzyme-catalyzed reaction (B) can go through several transition states. It is the transition state with the highest energy (more...)
A dramatic proof that stabilizing a transition state can greatly increase a reaction rate is provided by the intentional production of antibodies that act like enzymes. Consider, for example, the hydrolysis of an amide bond, which is similar to the peptide bond that joins two adjacent amino acids in a protein. In an aqueous solution, an amide bond hydrolyzes very slowly by the mechanism shown in Figure 3-47A. In the central intermediate, or transition state, the carbonyl carbon is bonded to four atoms arranged at the corners of a tetrahedron. By generating monoclonal antibodies that bind tightly to a stable analog of this very unstable tetrahedral intermediate, an antibody that functions like an enzyme can be obtained (Figure 3-47B). Because this catalytic antibody binds to and stabilizes the tetrahedral intermediate, it increases the spontaneous rate of amide-bond hydrolysis by more than 10,000-fold.
Catalytic antibodies. The stabilization of a transition state by an antibody creates an enzyme. (A) The reaction path for the hydrolysis of an amide bond goes through a tetrahedral intermediate, the high-energy transition state for the reaction. (B) The (more...)
Figure 3-48 compares the spontaneous reaction rates and the corresponding enzyme-catalyzed rates for five enzymes. Rate accelerations of 109–1023 are observed. Clearly, enzymes are much better catalysts than catalytic antibodies. Enzymes not only bind tightly to a transition state, they also contain precisely positioned atoms that alter the electron distributions in those atoms that participate directly in the making and breaking of covalent bonds. Peptide bonds, for example, can be hydrolyzed in the absence of an enzyme by exposing a polypeptide to either a strong acid or a strong base, as explained in Figure 3-49. Enzymes are unique, however, in being able to use acid and base catalysis simultaneously, since the acidic and basic residues required are prevented from combining with each other (as they would do in solution) by being tied to the rigid framework of the protein itself (Figure 3-49D).
The rate accelerations caused by five different enzymes. (Modified from A. Radzicka and R. Wolfenden, Science 267:90–93, 1995.)
Acid catalysis and base catalysis. (A) The start of the uncatalyzed reaction shown in Figure 3-47A is diagrammed, with blue indicating electron distribution in the water and carbonyl bonds. (B) An acid likes to donate a proton (H+) to other atoms. By (more...)
The fit between an enzyme and its substrate needs to be precise. A small change introduced by genetic engineering in the active site of an enzyme can have a profound effect. Replacing a glutamic acid with an aspartic acid in one enzyme, for example, shifts the position of the catalytic carboxylate ion by only 1 Å (about the radius of a hydrogen atom); yet this is enough to decrease the activity of the enzyme a thousandfold.
To demonstrate how enzymes catalyze chemical reactions, we shall use the example of an enzyme that acts as a natural antibiotic in egg white, saliva, tears, and other secretions. Lysozyme is an enzyme that catalyzes the cutting of polysaccharide chains in the cell walls of bacteria. Because the bacterial cell is under pressure from osmotic forces, cutting even a small number of polysaccharide chains causes the cell wall to rupture and the cell to burst. Lysozyme is a relatively small and stable protein that can be easily isolated in large quantities. For these reasons, it has been intensively studied, and it was the first enzyme to have its structure worked out in atomic detail by x-ray crystallography.
The reaction catalyzed by lysozyme is a hydrolysis: a molecule of water is added to a single bond between two adjacent sugar groups in the polysaccharide chain, thereby causing the bond to break (see Figure 2-19). The reaction is energetically favorable because the free energy of the severed polysaccharide chain is lower than the free energy of the intact chain. However, the pure polysaccharide can sit for years in water without being hydrolyzed to any detectable degree. This is because there is an energy barrier to the reaction, as discussed in Chapter 2 (see Figure 2-46). For a colliding water molecule to break a bond linking two sugars, the polysaccharide molecule has to be distorted into a particular shape—the transition state—in which the atoms around the bond have an altered geometry and electron distribution. Because of this distortion, a large activation energy must be supplied through random collisions before the reaction can take place. In an aqueous solution at room temperature, the energy of collisions almost never exceeds the activation energy. Consequently, hydrolysis occurs extremely slowly, if at all.
This situation is drastically changed when the polysaccharide binds to lysozyme. The active site of lysozyme, because its substrate is a polymer, is a long groove that holds six linked sugars at the same time. As soon as the polysaccharide binds to form an enzyme–substrate complex, the enzyme cuts the polysaccharide by adding a water molecule across one of its sugar–sugar bonds. The product chains are then quickly released, freeing the enzyme for further cycles of reaction (Figure 3-50).
The reaction catalyzed by lysozyme. (A) The enzyme lysozyme (denoted as E) catalyzes the cutting of a polysaccharide chain, which is its substrate (S). The enzyme first binds to the chain to form an enzyme–substrate complex (ES) and then catalyzes (more...)
The chemistry that underlies the binding of lysozyme to its substrate is the same as that for antibody binding to its antigen—the formation of multiple noncovalent bonds. However, lysozyme holds its polysaccharide substrate in a particular way, so that one of the two sugars involved in the bond to be broken is distorted from its normal, most stable conformation. The bond to be broken is also held close to two amino acids with acidic side chains (a glutamic acid and an aspartic acid) within the active site.
Conditions are thereby created in the microenvironment of the lysozyme active site that greatly reduce the activation energy necessary for the hydrolysis to take place. Figure 3-51 shows three central steps in this enzymatically catalyzed reaction.
Events at the active site of lysozyme. The top left and top right drawings depict the free substrate and the free products, respectively, whereas the other three drawings depict the sequential events at the enzyme active site. Note the change in the conformation (more...)
The enzyme stresses its bound substrate by bending some critical chemical bonds in one sugar, so that the shape of this sugar more closely resembles the shape of high-energy transition states formed during the reaction.
The negatively charged aspartic acid reacts with the C1 carbon atom on the distorted sugar, breaking the sugar–sugar bond and leaving the aspartic acid side chain covalently linked to the site of bond cleavage.
Aided by the negatively charged glutamic acid, a water molecule reacts with the C1 carbon atom, displacing the aspartic acid side chain and completing the process of hydrolysis.
The overall chemical reaction, from the initial binding of the polysaccharide on the surface of the enzyme through the final release of the severed chains, occurs many millions of times faster than it would in the absence of enzyme.
Similar mechanisms are used by other enzymes to lower activation energies and speed up the reactions they catalyze. In reactions involving two or more reactants, the active site also acts like a template, or mold, that brings the substrates together in the proper orientation for a reaction to occur between them (Figure 3-52A). As we saw for lysozyme, the active site of an enzyme contains precisely positioned atoms that speed up a reaction by using charged groups to alter the distribution of electrons in the substrates (Figure 3-52B). The binding to the enzyme also changes substrate shapes, bending bonds so as to drive a substrate toward a particular transition state (Figure 3-52C). Finally, like lysozyme, many enzymes participate intimately in the reaction by briefly forming a covalent bond between the substrate and a side chain of the enzyme. Subsequent steps in the reaction restore the side chain to its original state, so that the enzyme remains unchanged after the reaction (see also Figure 2-73).
Some general strategies of enzyme catalysis. (A) Holding substrates together in a precise alignment. (B) Charge stabilization of reaction intermediates. (C) Altering bond angles in the substrate to increase the rate of a particular reaction.
Although we have emphasized the versatility of proteins as chains of amino acids that perform different functions, there are many instances in which the amino acids by themselves are not enough. Just as humans employ tools to enhance and extend the capabilities of their hands, proteins often use small nonprotein molecules to perform functions that would be difficult or impossible to do with amino acids alone. Thus, the signal receptor protein rhodopsin, which is made by the photoreceptor cells in the retina, detects light by means of a small molecule, retinal, embedded in the protein (Figure 3-53A). Retinal changes its shape when it absorbs a photon of light, and this change causes the protein to trigger a cascade of enzymatic reactions that eventually leads to an electrical signal being carried to the brain.
Retinal and heme. (A) The structure of retinal, the light-sensitive molecule attached to rhodopsin in the eye. (B) The structure of a heme group. The carbon-containing heme ring is red and the iron atom at its center is orange. A heme group is tightly (more...)
Another example of a protein that contains a nonprotein portion is hemoglobin (see Figure 3-23). A molecule of hemoglobin carries four heme groups, ring-shaped molecules each with a single central iron atom (Figure 3-53B). Heme gives hemoglobin (and blood) its red color. By binding reversibly to oxygen gas through its iron atom, heme enables hemoglobin to pick up oxygen in the lungs and release it in the tissues.
Sometimes these small molecules are attached covalently and permanently to their protein, thereby becoming an integral part of the protein molecule itself. We see in Chapter 10 that proteins are often anchored to cell membranes through covalently attached lipid molecules. And membrane proteins exposed on the surface of the cell, as well as proteins secreted outside the cell, are often modified by the covalent addition of sugars and oligosaccharides.
Enzymes frequently have a small molecule or metal atom tightly associated with their active site that assists with their catalytic function. Carboxypeptidase, for example, an enzyme that cuts polypeptide chains, carries a tightly bound zinc ion in its active site. During the cleavage of a peptide bond by carboxypeptidase, the zinc ion forms a transient bond with one of the substrate atoms, thereby assisting the hydrolysis reaction. In other enzymes, a small organic molecule serves a similar purpose. Such organic molecules are often referred to as coenzymes. An example is biotin, which is found in enzymes that transfer a carboxylate group (–COO–) from one molecule to another (see Figure 2-63). Biotin participates in these reactions by forming a transient covalent bond to the –COO– group to be transferred, being better suited to this function than any of the amino acids used to make proteins. Because it cannot be synthesized by humans, and must therefore be supplied in small quantities in our diet, biotin is a vitamin, as are many other coenzymes (Table 3-2). Other vitamins are needed to make other small molecules that are essential components of our proteins; vitamin A, for example, is needed in the diet to make retinal, the light-sensitive part of rhodopsin.
Many Vitamins Provide Critical Coenzymes for Human Cells.
The efficiency of enzymes in accelerating chemical reactions is crucial to the maintenance of life. Cells, in effect, must race against the unavoidable processes of decay, which—if left unattended—cause macromolecules to run downhill toward greater and greater disorder. If the rates of desirable reactions were not greater than the rates of competing side reactions, a cell would soon die. Some idea of the rate at which cell metabolism proceeds can be obtained by measuring the rate of ATP utilization. A typical mammalian cell “turns over” (i.e., hydrolyzes and restores by phosphorylation) its entire ATP pool once every 1 or 2 minutes. For each cell this turnover represents the utilization of roughly 107 molecules of ATP per second (or, for the human body, about 1 gram of ATP every minute).
The rates of reactions in cells are rapid because of the effectiveness of enzyme catalysis. Many important enzymes have become so efficient that there is no possibility of further useful improvement. The factor that limits the reaction rate is no longer the enzyme’s intrinsic speed of action; rather, it is the frequency with which the enzyme collides with its substrate. Such a reaction is said to be diffusion-limited.
If an enzyme-catalyzed reaction is diffusion-limited, its rate depends on the concentration of both the enzyme and its substrate. If a sequence of reactions is to occur extremely rapidly, each metabolic intermediate and enzyme involved must be present in high concentration. However, given the enormous number of different reactions performed by a cell, there are limits to the concentrations of substrates that can be achieved. In fact, most metabolites are present in micromolar (10–6 M) concentrations, and most enzyme concentrations are much lower. How is it possible, therefore, to maintain very fast metabolic rates?
The answer lies in the spatial organization of cell components. Reaction rates can be increased without raising substrate concentrations by bringing the various enzymes involved in a reaction sequence together to form a large protein assembly known as a multienzyme complex (Figure 3-54). Because this allows the product of enzyme A to be passed directly to enzyme B, and so on, diffusion rates need not be limiting, even when the concentrations of the substrates in the cell as a whole are very low. It is perhaps not surprising, therefore, that such enzyme complexes are very common, and they are involved in nearly all aspects of metabolism—including the central genetic processes of DNA, RNA, and protein synthesis. In fact, few enzymes in eucaryotic cells may be left to diffuse freely in solution; instead, most seem to have evolved binding sites that concentrate them with other proteins of related function in particular regions of the cell, thereby increasing the rate and efficiency of the reactions that they catalyze.
The structure of pyruvate dehydrogenase. This enzyme complex catalyzes the conversion of pyruvate to acetyl CoA, as part of the pathway that oxidizes sugars to CO2 and H2O. It is an example of a large multienzyme complex in which reaction intermediates (more...)
Eucaryotic cells have yet another way of increasing the rate of metabolic reactions, using their intracellular membrane systems. These membranes can segregate particular substrates and the enzymes that act on them into the same membrane-enclosed compartment, such as the endoplasmic reticulum or the cell nucleus. If, for example, a compartment occupies a total of 10% of the volume of the cell, the concentration of reactants in the compartment may be increased as much as 10 times compared with the same cell with no compartmentalization. Reactions that would otherwise be limited by the speed of diffusion can thereby be speeded up by a factor of 10.
A living cell contains thousands of enzymes, many of which operate at the same time and in the same small volume of the cytosol. By their catalytic action, these enzymes generate a complex web of metabolic pathways, each composed of chains of chemical reactions in which the product of one enzyme becomes the substrate of the next. In this maze of pathways, there are many branch points where different enzymes compete for the same substrate. The system is so complex (see Figure 2-88) that elaborate controls are required to regulate when and how rapidly each reaction occurs.
Regulation occurs at many levels. At one level, the cell controls how many molecules of each enzyme it makes by regulating the expression of the gene that encodes that enzyme (discussed in Chapter 7). The cell also controls enzymatic activities by confining sets of enzymes to particular subcellular compartments, enclosed by distinct membranes (discussed in Chapters 12 and 14). The rate of protein destruction by targeted proteolysis represents yet another important regulatory mechanism (see p. 361). But the most rapid and general process that adjusts reaction rates operates through a direct, reversible change in the activity of an enzyme in response to specific molecules that it encounters.
The most common type of control occurs when a molecule other than one of the substrates binds to an enzyme at a special regulatory site outside the active site, thereby altering the rate at which the enzyme converts its substrates to products. In feedback inhibition, an enzyme acting early in a reaction pathway is inhibited by a late product of that pathway. Thus, whenever large quantities of the final product begin to accumulate, this product binds to the first enzyme and slows down its catalytic action, thereby limiting the further entry of substrates into that reaction pathway (Figure 3-55). Where pathways branch or intersect, there are usually multiple points of control by different final products, each of which works to regulate its own synthesis (Figure 3-56). Feedback inhibition can work almost instantaneously and is rapidly reversed when the level of the product falls.
Feedback inhibition of a single biosynthetic pathway. The end-product Z inhibits the first enzyme that is unique to its synthesis and thereby controls its own level in the cell. This is an example of negative regulation.
Multiple feedback inhibition. In this example, which shows the biosynthetic pathways for four different amino acids in bacteria, the red arrows indicate positions at which products feed back to inhibit enzymes. Each amino acid controls the first enzyme (more...)
Feedback inhibition is negative regulation: it prevents an enzyme from acting. Enzymes can also be subject to positive regulation, in which the enzyme’s activity is stimulated by a regulatory molecule rather than being shut down. Positive regulation occurs when a product in one branch of the metabolic maze stimulates the activity of an enzyme in another pathway. As one example, the accumulation of ADP activates several enzymes involved in the oxidation of sugar molecules, thereby stimulating the cell to convert more ADP to ATP.
One feature of feedback inhibition was initially puzzling to those who discovered it: the regulatory molecule often has a shape totally different from the shape of the substrate of the enzyme. This is why this form of regulation is termed allostery (from the Greek words allos, meaning “other,” and stereos, meaning “solid” or “three-dimensional”). As more was learned about feedback inhibition, it was recognized that many enzymes must have at least two different binding sites on their surface—an active site that recognizes the substrates, and a regulatory site that recognizes a regulatory molecule. These two sites must somehow communicate in a way that allows the catalytic events at the active site to be influenced by the binding of the regulatory molecule at its separate site on the protein’s surface.
The interaction between separated sites on a protein molecule is now known to depend on a conformational change in the protein: binding at one of the sites causes a shift from one folded shape to a slightly different folded shape. During feedback inhibition, for example, the binding of an inhibitor at one site on the protein causes the protein to shift to a conformation in which its active site—located elsewhere in the protein—becomes incapacitated.
It is thought that most protein molecules are allosteric. They can adopt two or more slightly different conformations, and a shift from one to another caused by the binding of a ligand can alter their activity. This is true not only for enzymes but also for many other proteins—including receptors, structural proteins, and motor proteins. There is nothing mysterious about the allosteric regulation of these proteins: each conformation of the protein has somewhat different surface contours, and the protein’s binding sites for ligands are altered when the protein changes shape. Moreover as we discuss next, each ligand stabilizes the conformation that it binds to most strongly, and thus—at high enough concentrations—tends to “switch” the protein toward the conformation it prefers.
The effects of ligand binding on a protein follow from a fundamental chemical principle known as linkage. Suppose, for example, that a protein that binds glucose also binds another molecule, X, at a distant site on the protein’s surface. If the binding site for X changes shape as part of the conformational change induced by glucose binding, the binding sites for X and for glucose are said to be coupled. Whenever two ligands prefer to bind to the same conformation of an allosteric protein, it follows from basic thermodynamic principles that each ligand must increase the affinity of the protein for the other. Thus, if the shift of the protein in Figure 3-57 to the closed conformation that binds glucose best also causes the binding site for X to fit X better, then the protein will bind glucose more tightly when X is present than when X is absent.
Positive regulation caused by conformational coupling between two distant binding sites. In this example, both glucose and molecule X bind best to the closed conformation of a protein with two domains. Because both glucose and molecule X drive the protein (more...)
Conversely, linkage operates in a negative way if two ligands prefer to bind to different conformations of the same protein. In this case, the binding of the first ligand discourages the binding of the second ligand. Thus, if a shape change caused by glucose binding decreases the affinity of a protein for molecule X, the binding of X must also decrease the protein’s affinity for glucose (Figure 3-58). The linkage relationship is quantitatively reciprocal, so that, for example, if glucose has a very large effect on the binding of X, X has a very large effect on the binding of glucose.
Negative regulation caused by conformational coupling between two distant binding sites. The scheme here resembles that in the previous figure, but here molecule X prefers the open conformation, while glucose prefers the closed conformation. Because glucose (more...)
The relationships shown in Figures 3-57 and 3-58 underlie all of cell biology. They seem so obvious in retrospect that we now take them for granted. But their discovery in the 1950s, followed by a general description of allostery in the early 1960s, was revolutionary at the time. Since molecule X in these examples binds at a site that is distinct from the site where catalysis occurs, it need have no chemical relationship to glucose or to any other ligand that binds at the active site. As we have just seen, for enzymes that are regulated in this way, molecule X can either turn the enzyme on (positive regulation) or turn it off (negative regulation). By such a mechanism, allosteric proteins serve as general switches that, in principle, allow one molecule in a cell to affect the fate of any other.
A single subunit enzyme that is regulated by negative feedback can at most decrease from 90% to about 10% activity in response to a 100-fold increase in the concentration of an inhibitory ligand that it binds (Figure 3-59, red line). Responses of this type are apparently not sharp enough for optimal cell regulation, and most enzymes that are turned on or off by ligand binding consist of symmetric assemblies of identical subunits. With this arrangement, the binding of a molecule of ligand to a single site on one subunit can trigger an allosteric change in the subunit that can be transmitted to the neighboring subunits, helping them to bind the same ligand. As a result, a cooperative allosteric transition occurs (Figure 3-59, blue line), allowing a relatively small change in ligand concentration in the cell to switch the whole assembly from an almost fully active to an almost fully inactive conformation (or vice versa).
Enzyme activity versus the concentration of inhibitory ligand for single-subunit and multi-subunit allosteric enzymes. For an enzyme with a single subunit (red line), a drop from 90% enzyme activity to 10% activity (indicated by the two dots on the curve) (more...)
The principles involved in a cooperative “all-or-none” transition are easiest to visualize for an enzyme that forms a symmetric dimer. In the example shown in Figure 3-60, the first molecule of an inhibitory ligand binds with great difficulty since its binding destroys an energetically favorable interaction between the two identical monomers in the dimer. A second molecule of inhibitory ligand now binds more easily, however, because its binding restores the monomer–monomer contacts of a symmetric dimer (and also completely inactivates the enzyme).
A cooperative allosteric transition in an enzyme composed of two identical subunits. This diagram illustrates how the conformation of one subunit can influence that of its neighbor. The binding of a single molecule of an inhibitory ligand (yellow) to (more...)
An even sharper response to a ligand can be obtained with larger assemblies, such as the enzyme formed from 12 polypeptide chains that we discuss next.
One enzyme used in the early studies of allosteric regulation was aspartate transcarbamoylase from E. coli. It catalyzes the important reaction that begins the synthesis of the pyrimidine ring of C, U, and T nucleotides: carbamoylphosphate + aspartate → N-carbamoylaspartate. One of the final products of this pathway, cytosine triphosphate (CTP), binds to the enzyme to turn it off whenever CTP is plentiful.
Aspartate transcarbamoylase is a large complex of six regulatory and six catalytic subunits. The catalytic subunits are present as two trimers, each arranged like an equilateral triangle; the two trimers face each other and are held together by three regulatory dimers that form a bridge between them. The entire molecule is poised to undergo a concerted, all-or-none, allosteric transition between two conformations, designated as T (tense) and R (relaxed) states (Figure 3-61).
The transition between R and T states in the enzyme aspartate transcarbamoylase. The enzyme consists of a complex of six catalytic subunits and six regulatory subunits, and the structures of its inactive (T state) and active (R state) forms have been (more...)
The binding of substrates (carbamoylphosphate and aspartate) to the catalytic trimers drives aspartate transcarbamoylase into its catalytically active R state, from which the regulatory CTP molecules dissociate. By contrast, the binding of CTP to the regulatory dimers converts the enzyme to the inactive T state, from which the substrates dissociate. This tug-of-war between CTP and substrates is identical in principle to that described previously in Figure 3-58 for a simpler allosteric protein. But because the tug-of-war occurs in a symmetric molecule with multiple binding sites, the enzyme undergoes a cooperative allosteric transition that will turn it on suddenly as substrates accumulate (forming the R state) or shut it off rapidly when CTP accumulates (forming the T state).
A combination of biochemistry and x-ray crystallography has revealed many fascinating details of this allosteric transition. Each regulatory subunit has two domains, and the binding of CTP causes the two domains to move relative to each other, so that they function like a lever that rotates the two catalytic trimers and pulls them closer together into the T state (see Figure 3-61). When this occurs, hydrogen bonds form between opposing catalytic subunits that help widen the cleft that forms the active site within each catalytic subunit, thereby destroying the binding sites for the substrates (Figure 3-62). Adding large amounts of substrate has the opposite effect, favoring the R state by binding in the cleft of each catalytic subunit and opposing the above conformational change. Conformations that are intermediate between R and T are unstable, so that the enzyme mostly clicks back and forth between its R and T forms, producing a mixture of these two species in proportions that depend on the relative concentrations of CTP and substrates.
Part of the on–off switch in the catalytic subunits of aspartate transcarbamoylase. Changes in the indicated hydrogen-bonding interactions are partly responsible for switching this enzyme’s active site between active (yellow) and inactive (more...)
Enzymes are regulated by more than the binding of small molecules. A second method that is commonly used by eucaryotic cells to regulate a protein’s function is the covalent addition of a phosphate group to one of its amino acid side chains. Such phosphorylation events can affect the protein in two important ways.
First, because each phosphate group carries two negative charges, the enzyme-catalyzed addition of a phosphate group to a protein can cause a major conformational change in the protein by, for example, attracting a cluster of positively charged amino acid side chains. This can, in turn, affect the binding of ligands elsewhere on the protein surface, dramatically changing the protein’s activity through an allosteric effect. Removal of the phosphate group by a second enzyme returns the protein to its original conformation and restores its initial activity.
Second, an attached phosphate group can form part of a structure that is directly recognized by binding sites of other proteins. As previously discussed, certain small protein domains, called modules, appear very frequently in larger proteins. A large number of these modules provide binding sites for attaching their protein to phosphorylated peptides in other protein molecules. One of these modules is the SH2 domain, featured previously in this chapter, which binds to a short peptide sequence containing a phosphorylated tyrosine side chain (see Figure 3-40B). Several other types of modules recognize phosphorylated serine or threonine side chains in a specific context. As a result, protein phosphorylation and dephosphorylation events have a major role in driving the regulated assembly and disassembly of protein complexes.
Through such effects, reversible protein phosphorylation controls the activity structure and cellular localization of many types of proteins in eucaryotic cells. In fact, this regulation is so extensive that more than one-third of the 10,000 or so proteins in a typical mammalian cell are thought to be phosphorylated at any given time—many with more than one phosphate. As might be expected, the addition and removal of phosphate groups from specific proteins often occur in response to signals that specify some change in a cell’s state. For example, the complicated series of events that takes place as a eucaryotic cell divides is largely timed in this way (discussed in Chapter 17), and many of the signals mediating cell–cell interactions are relayed from the plasma membrane to the nucleus by a cascade of protein phosphorylation events (discussed in Chapter 15).
Protein phosphorylation involves the enzyme-catalyzed transfer of the terminal phosphate group of an ATP molecule to the hydroxyl group on a serine, threonine, or tyrosine side chain of the protein (Figure 3-63). This reaction is catalyzed by a protein kinase, and the reaction is essentially unidirectional because of the large amount of free energy released when the phosphate–phosphate bond in ATP is broken to produce ADP (discussed in Chapter 2). The reverse reaction of phosphate removal, or dephosphorylation, is instead catalyzed by a protein phosphatase. Cells contain hundreds of different protein kinases, each responsible for phosphorylating a different protein or set of proteins. There are also many different protein phosphatases; some of these are highly specific and remove phosphate groups from only one or a few proteins, whereas others act on a broad range of proteins and are targeted to specific substrates by regulatory subunits. The state of phosphorylation of a protein at any moment, and thus its activity, depends on the relative activities of the protein kinases and phosphatases that modify it.
Protein phosphorylation. Many thousands of proteins in a typical eucaryotic cell are modified by the covalent addition of a phosphate group. (A) The general reaction, shown here, entails the transfer of a phosphate group from ATP to an amino acid side (more...)
The protein kinases that phosphorylate proteins in eucaryotic cells belong to a very large family of enzymes, which share a catalytic (kinase) sequence of 250 amino acids. The various family members contain different amino acid sequences on either side of the kinase sequence (see Figure 3-12), and often have short amino acid sequences inserted into loops within it (red arrowheads in Figure 3-64). Some of these additional amino acid sequences enable each kinase to recognize the specific set of proteins it phosphorylates, or to bind to structures that localize it in specific regions of the cell. Other parts of the protein allow the activity of each enzyme to be tightly regulated, so it can be turned on and off in response to different specific signals, as described below.
The three-dimensional structure of a protein kinase. Superimposed on this structure are red arrowheads to indicate sites where insertions of 5–100 amino acids are found in some members of the protein kinase family. These insertions are located (more...)
By comparing the number of amino acid sequence differences between the various members of a protein family, one can construct an “evolutionary tree” that is thought to reflect the pattern of gene duplication and divergence that gave rise to the family. An evolutionary tree of protein kinases is shown in Figure 3-65. Not surprisingly, kinases with related functions are often located on nearby branches of the tree: the protein kinases involved in cell signaling that phosphorylate tyrosine side chains, for example, are all clustered in the top left corner of the tree. The other kinases shown phosphorylate either a serine or a threonine side chain, and many are organized into clusters that seem to reflect their function—in transmembrane signal transduction, intracellular signal amplification, cell-cycle control, and so on.
An evolutionary tree of selected protein kinases. Although a higher eucaryotic cell contains hundreds of such enzymes, and the human genome codes for more than 500, only some of those discussed in this book are shown.
As a result of the combined activities of protein kinases and protein phosphatases, the phosphate groups on proteins are continually turning over—being added and then rapidly removed. Such phosphorylation cycles may seem wasteful, but they are important in allowing the phosphorylated proteins to switch rapidly from one state to another: the more rapid the cycle, the faster the state of phosphorylation of a population of protein molecules can change in response to a sudden stimulus that changes the phosphorylation rate (see Figure 15-10). The energy required to drive the phosphorylation cycle is derived from the free energy of ATP hydrolysis, one molecule of which is consumed for each phosphorylation event.
The hundreds of different protein kinases in a eucaryotic cell are organized into complex networks of signaling pathways that help to coordinate the cell’s activities, drive the cell cycle, and relay signals into the cell from the cell’s environment. Many of the extracellular signals involved need to be both integrated and amplified by the cell. Individual protein kinases (and other signaling proteins) serve as input–output devices, or “microchips,” in the integration process. An important part of the input to these proteins comes from the control that is exerted by phosphates added and removed from them by protein kinases and protein phosphatases, respectively, in the signaling network.
For a protein that is phosphorylated at multiple sites, specific sets of phosphate groups serve to activate the protein, while other sets can inactivate it. A cyclin-dependent protein kinase (Cdk) represents a good example of such a signal processing device. Kinases in this class phosphorylate serines, and they are central components of the cell-cycle control system in eucaryotic cells, as discussed in detail in Chapter 17. In a vertebrate cell, individual Cdk enzymes turn on and off in succession, as a cell proceeds through the different phases of its division cycle. When a particular one of these kinases is on, it influences various aspects of cell behavior through effects on the proteins it phosphorylates.
A Cdk protein becomes active as a serine/threonine protein kinase only when it is bound to a second protein called a cyclin. But, as Figure 3-66 shows, the binding of cyclin is only one of three distinct “inputs” required to activate the Cdk. In addition to cyclin binding, a phosphate must be added to a specific threonine side chain, and a phosphate elsewhere in the protein (covalently bound to a specific tyrosine side chain) must be removed. Cdk thus monitors a specific set of cell components—a cyclin, a protein kinase, and a protein phosphatase—and it acts as an input–output device that turns on if, and only if, each of these components has attained its appropriate activity state. Some cyclins rise and fall in concentration in step with the cell cycle, increasing gradually in amount until they are suddenly destroyed at a particular point in the cycle. The sudden destruction of a cyclin (by targeted proteolysis) immediately shuts off its partner Cdk enzyme, and this triggers a specific step in the cell cycle.
How a Cdk protein acts as an integrating device. The function of these central regulators of the cell cycle is discussed in Chapter 17.
A similar type of microchip behavior is exhibited by the Src family of protein kinases. The Src protein (pronounced “sarc”) was the first tyrosine kinase to be discovered, and it is now known to be part of a subfamily of nine very similar protein kinases, which are found only in multicellular animals. As indicated by the evolutionary tree in Figure 3-65, sequence comparisons suggest that tyrosine kinases as a group were a relatively late innovation that branched off from the serine/threonine kinases, with the Src subfamily being only one subgroup of the tyrosine kinases created in this way.
The Src protein and its homologs contain a short N-terminal region that becomes covalently linked to a strongly hydrophobic fatty acid, which holds the kinase at the cytoplasmic face of the plasma membrane. Next come two peptide-binding modules, a Src homology 3 (SH3) domain and a SH2 domain, followed by the kinase catalytic domains (Figure 3-67). These kinases normally exist in an inactive conformation, in which a phosphorylated tyrosine near the C-terminus is bound to the SH2 domain, and the SH3 domain is bound to an internal peptide in a way that distorts the active site of the enzyme and helps to render it inactive (see Figure 3-12).
The domain structure of the Src family of protein kinases, mapped along the amino acid sequence.
Turning the kinase on involves at least two specific inputs: removal of the C-terminal phosphate and the binding of the SH3 domain by a specific activating protein (Figure 3-68). As for the Cdk protein, the activation of the Src kinase signals that a particular set of separate upstream events has been completed (Figure 3-69). Thus, both the Cdk and Src families of proteins serve as specific signal integrators, helping to generate the complex web of information-processing events that enable the cell to compute logical responses to a complex set of conditions.
The activation of a Src-type protein kinase by two sequential events. (Adapted from I. Moareti, et al., Nature 385:650–653, 1997.)
How a Src-type protein kinase acts as an integrating device. The disruption of the SH3 domain interaction (green) can involve replacing its binding to the indicated red linker region with a tighter interaction with the Nef protein, as illustrated in Figure (more...)
We have described how the addition or removal of phosphate groups on a protein can be used by a cell to control the protein’s activity. In the examples discussed so far, the phosphate is transferred from an ATP molecule to an amino acid side chain of the protein in a reaction catalyzed by a specific protein kinase. Eucaryotic cells also have another way to control protein activity by phosphate addition and removal. In this case, the phosphate is not attached directly to the protein; instead, it is a part of the guanine nucleotide GTP, which binds very tightly to the protein. In general, proteins regulated in this way are in their active conformations with GTP bound. The loss of a phosphate group occurs when the bound GTP is hydrolyzed to GDP in a reaction catalyzed by the protein itself, and in its GDP bound state the protein is inactive. In this way, GTP-binding proteins act as on–off switches whose activity is determined by the presence or absence of an additional phosphate on a bound GDP molecule (Figure 3-70).
GTP-binding proteins as molecular switches. The activity of a GTP-binding protein (also called a GTPase) generally requires the presence of a tightly bound GTP molecule (switch “on”). Hydrolysis of this GTP molecule produces GDP and inorganic (more...)
GTP-binding proteins (also called GTPases because of the GTP hydrolysis they catalyze) constitute a large family of proteins that all contain variations on the same GTP-binding globular domain. When the tightly bound GTP is hydrolyzed to GDP, this domain undergoes a conformational change that inactivates it. The three-dimensional structure of a prototypical member of this family, the monomeric GTPase called Ras, is shown in Figure 3-71.
The structure of the Ras protein in its GTP-bound form. This monomeric GTPase illustrates the structure of a GTP-binding domain, which is present in a large family of GTP-binding proteins. The red regions change their conformation when the GTP molecule (more...)
The Ras protein has an important role in cell signaling (discussed in Chapter 15). In its GTP-bound form, it is active and stimulates a cascade of protein phosphorylations in the cell. Most of the time, however, the protein is in its inactive, GDP-bound form. It becomes active when it exchanges its GDP for a GTP molecule in response to extracellular signals, such as growth factors, that bind to receptors in the plasma membrane (see Figure 15-55).
GTP-binding proteins are controlled by regulatory proteins that determine whether GTP or GDP is bound, just as phosphorylated proteins are turned on and off by protein kinases and protein phosphatases. Thus, Ras is inactivated by a GTPase-activating protein (GAP), which binds to the Ras protein and induces it to hydrolyze its bound GTP molecule to GDP—which remains tightly bound—and inorganic phosphate (Pi)—which is rapidly released. The Ras protein stays in its inactive, GDP-bound conformation until it encounters a guanine nucleotide exchange factor (GEF), which binds to GDP-Ras and causes it to release its GDP. Because the empty nucleotide-binding site is immediately filled by a GTP molecule (GTP is present in large excess over GDP in cells), the GEF activates Ras by indirectly adding back the phosphate removed by GTP hydrolysis. Thus, in a sense, the roles of GAP and GEF are analogous to those of a protein phosphatase and a protein kinase, respectively (Figure 3-72).
A comparison of the two major intracellular signaling mechanisms in eucaryotic cells. In both cases, a signaling protein is activated by the addition of a phosphate group and inactivated by the removal of this phosphate. To emphasize the similarities (more...)
The Ras protein belongs to a large superfamily of monomeric GTPases, each of which consists of a single GTP-binding domain of about 200 amino acids. Over the course of evolution, this domain has also become joined to larger proteins with additional domains, creating a large family of GTP-binding proteins. Family members include the receptor-associated trimeric G proteins involved in cell signaling (discussed in Chapter 15), proteins regulating the traffic of vesicles between intracellular compartments (discussed in Chapter 13), and proteins that bind to transfer RNA and are required as assembly factors for protein synthesis on the ribosome (discussed in Chapter 6). In each case, an important biological activity is controlled by a change in the protein’s conformation that is caused by GTP hydrolysis in a Ras-like domain.
The EF-Tu protein provides a good example of how this family of proteins works. EF-Tu is an abundant molecule that serves as an elongation factor (hence the EF) in protein synthesis, loading each amino-acyl tRNA molecule onto the ribosome. The tRNA molecule forms a tight complex with the GTP-bound form of EF-Tu (Figure 3-73). In this complex, the amino acid attached to the tRNA is masked. Unmasking is required for the tRNA to transfer its amino acid in protein synthesis, and it occurs on the ribosome when the GTP bound to EF-Tu is hydrolyzed, allowing the tRNA to dissociate. Since the GTP hydrolysis is triggered by a proper fit of the tRNA to the mRNA molecule on the ribosome, the EF-Tu serves as an assembly factor that discriminates between correct and incorrect mRNA–tRNA pairings (see Figure 6-66 for a further discussion of this function of EF-Tu).
An aminoacyl tRNA molecule bound to EF-Tu. The three domains of the EF-Tu protein are colored differently, to match Figure 3-74. This is a bacterial protein; however, a very similar protein exists in eukaryotes, where it is called EF-1. (Coordinates determined (more...)
Comparison of the three-dimensional structure of EF-Tu in its GTP-bound and GDP-bound forms reveals how the unmasking of the tRNA occurs. The dissociation of the inorganic phosphate group (Pi), which follows the reaction GTP → GDP + Pi, causes a shift of a few tenths of a nanometer at the GTP-binding site, just as it does in the Ras protein. This tiny movement, equivalent to a few times the diameter of a hydrogen atom, causes a conformational change to propagate along a crucial piece of α helix, called the switch helix, in the Ras-like domain of the protein. The switch helix seems to serve as a latch that adheres to a specific site in another domain of the molecule, holding the protein in a “shut” conformation. The conformational change triggered by GTP hydrolysis causes the switch helix to detach, allowing separate domains of the protein to swing apart, through a distance of about 4 nm. This releases the bound tRNA molecule, allowing its attached amino acid to be used (Figure 3-74).
The large conformational change in EF-Tu caused by GTP hydrolysis. (A) The three-dimensional structure of EF-Tu with GTP bound. The domain at the top is homologous to the Ras protein, and its red α helix is the switch helix, which moves after (more...)
One can see from this example how cells have exploited a simple chemical change that occurs on the surface of a small protein domain to create a movement 50 times larger. Dramatic shape changes of this type also underlie the very large movements that occur in motor proteins, as we discuss next.
We have seen how conformational changes in proteins have a central role in enzyme regulation and cell signaling. We now discuss proteins whose major function is to move other molecules. These motor proteins generate the forces responsible for muscle contraction and the crawling and swimming of cells. Motor proteins also power smaller-scale intracellular movements: they help to move chromosomes to opposite ends of the cell during mitosis (discussed in Chapter 18), to move organelles along molecular tracks within the cell (discussed in Chapter 16), and to move enzymes along a DNA strand during the synthesis of a new DNA molecule (discussed in Chapter 5). All these fundamental processes depend on proteins with moving parts that operate as force-generating machines.
How do these machines work? In other words, how are shape changes in proteins used to generate directed movements in cells? If, for example, a protein is required to walk along a narrow thread such as a DNA molecule, it can do this by undergoing a series of conformational changes, such as those shown in Figure 3-75. With nothing to drive these changes in an orderly sequence, however, they are perfectly reversible, and the protein can wander randomly back and forth along the thread. We can look at this situation in another way. Since the directional movement of a protein does work, the laws of thermodynamics (discussed in Chapter 2) demand that such movement utilize free energy from some other source (otherwise the protein could be used to make a perpetual motion machine). Therefore, without an input of energy, the protein molecule can only wander aimlessly.
An allosteric “walking” protein. Although its three different conformations allow it to wander randomly back and forth while bound to a thread or a filament, the protein cannot move uniformly in a single direction.
How, then, can one make the series of conformational changes unidirectional? To force the entire cycle to proceed in one direction, it is enough to make any one of the changes in shape irreversible. For most proteins that are able to walk in one direction for long distances, this is achieved by coupling one of the conformational changes to the hydrolysis of an ATP molecule bound to the protein. The mechanism is similar to the one just discussed that drives allosteric protein shape changes by GTP hydrolysis. Because a great deal of free energy is released when ATP (or GTP) is hydrolyzed, it is very unlikely that the nucleotide-binding protein will undergo the reverse shape change needed for moving backward—since this would require that it also reverse the ATP hydrolysis by adding a phosphate molecule to ADP to form ATP.
In the model shown in Figure 3-76, ATP binding shifts a motor protein from conformation 1 to conformation 2. The bound ATP is then hydrolyzed to produce ADP and inorganic phosphate (Pi), causing a change from conformation 2 to conformation 3. Finally, the release of the bound ADP and Pi drives the protein back to conformation 1. Because the transition 2 → 3 is driven by the energy provided by ATP hydrolysis, this series of conformational changes is effectively irreversible. Thus, the entire cycle goes in only one direction, causing the protein molecule to walk continuously to the right in this example.
An allosteric motor protein. The transition between three different conformations includes a step driven by the hydrolysis of a bound ATP molecule, and this makes the entire cycle essentially irreversible. By repeated cycles, the protein therefore moves (more...)
Many motor proteins generate directional movement in this general way, including the muscle motor protein myosin, which walks along actin filaments to generate muscle contraction, and the kinesin proteins that walk along microtubules (both discussed in Chapter 16). These movements can be rapid: some of the motor proteins involved in DNA replication (the DNA helicases) propel themselves along a DNA strand at rates as high as 1000 nucleotides per second.
We have thus far seen how allosteric proteins can act as microchips (Cdk and Src kinases), as assembly factors (EF-Tu), and as generators of mechanical force and motion (motor proteins). Allosteric proteins can also harness energy derived from ATP hydrolysis, ion gradients, or electron transport processes to pump specific ions or small molecules across a membrane. We consider one example here; others will be discussed in Chapter 11.
One of the best understood pump proteins is the calcium transport protein from muscle cells. This protein, called the Ca 2+ pump, is embedded in the membrane of a specialized organelle in muscle cells called the sarcoplasmic reticulum. The Ca2+ pump (also known as the Ca2+ ATPase) maintains the low cytoplasmic calcium concentration of resting muscle by pumping calcium out of the cytosol into the membrane-enclosed sarcoplasmic reticulum; then, in response to a nerve impulse, Ca2+ is rapidly released (through other channels) back into the cytosol to trigger muscle contraction. The Ca2+ pump is homologous to the Na+-K+ pump that maintains Na+ and K+ concentration differences across the plasma membrane, both being members of a family of P-type cation pumps—so named because these proteins are autophosphorylated during their reaction cycle. A large, cytoplasmic head region binds and hydrolyzes ATP, forming a covalent bond with the phosphate released. The protein thereby transiently shifts to a high-energy, phosphorylated state that is tightly bound to two Ca2+ ions that were picked up from the cytosol. This form of the protein then decays to a low-energy, phosphorylated state, which causes the Ca2+ ions to be released into the lumen. Dephosphorylation of the enzyme finally releases the phosphate and resets the protein for its next round of ion pumping.
The head region of all P-type cation pumps is linked to a series of ten transmembrane α helices, four of which form transmembrane Ca2+-binding sites for the Ca2+ pump. The three-dimensional structure of this protein has been deciphered by high-resolution electron microscopy and x-ray diffraction (see Figure 11-15). This has enabled biologists to derive a molecular model for pump action based on extensive biochemical data on its normal and mutant forms.
As illustrated in Figure 3-77A, the transmembrane α helices that bind the Ca2+ wind around each other and create a cavity for Ca2+ between the helices. ATP hydrolysis generates a series of major conformational changes in the cytoplasmic head, which—through its stalk connection to the transmembrane helices—changes the structure and relative orientations of some of the helices in the membrane, thereby altering the cavity in a way that pushes the Ca2+ ions unidirectionally across the membrane (Figure 3-77B). As for a motor protein, unidirectional transport of an ion requires the cycle to use energy, so as to impart a preferred directionality to the protein’s conformational changes.
The transport of calcium ions by the Ca2+ pump. (A) The structure of the pump protein, formed from a single subunit of 994 amino acids (see Figure 11-15 for details). (B) A model for ion pumping. For simplicity, only two of the four transmembrane α (more...)
Humans have invented many different types of mechanical pumps, and it should not be surprising that cells also contain membrane-bound pumps that function in other ways. Most notable are the rotary pumps that couple the hydrolysis of ATP to the transport of H+ ions (protons). These pumps resemble miniature turbines, and they are used to acidify the interior of lysosomes and other eucaryotic organelles. Like other ion pumps that create ion gradients, they can function in reverse to catalyze the reaction ADP + Pi → ATP if there is a steep enough gradient across their membrane of the ion that they transport.
One such pump, the ATP synthase, harnesses a gradient of proton concentration produced by electron transport processes to produce most of the ATP used in the living world. Because this ubiquitous pump has such a central role in energy conversion, we postpone discussing its three-dimensional structure and mechanism until Chapter 14.
As one progresses from small, single-domain proteins to large proteins formed from many domains, the functions the proteins can perform become more elaborate. The most impressive tasks, however, are carried out by large protein assemblies formed from many protein molecules. Now that it is possible to reconstruct most biological processes in cell-free systems in the laboratory, it is clear that each of the central processes in a cell—such as DNA replication, protein synthesis, vesicle budding, or transmembrane signaling—is catalyzed by a highly coordinated, linked set of ten or more proteins. In most such protein machines, the hydrolysis of bound nucleoside triphosphates (ATP or GTP) drives an ordered series of conformational changes in some of the individual protein subunits, enabling the ensemble of proteins to move coordinately. In this way, each of the appropriate enzymes can be placed directly into the positions where they are needed to perform successive reactions in a series. This is what occurs, for example, in protein synthesis on a ribosome (discussed in Chapter 6)—or in DNA replication, where a large multiprotein complex moves rapidly along the DNA (discussed in Chapter 5).
Cells have evolved protein machines for the same reason that humans have invented mechanical and electronic machines. For accomplishing almost any task, manipulations that are spatially and temporally coordinated through linked processes are much more efficient than the sequential use of individual tools.
There are many challenges facing cell biologists in this “post-genome” era when complete genome sequences are known. One is the need to dissect and reconstruct each one of the thousands of protein machines that exist in an organism such as ourselves. To understand these remarkable protein complexes, each must be reconstituted from its purified protein parts—so that its detailed mode of operation can be studied under controlled conditions in a test tube, free from all other cell components. This alone is a massive task. But we now know that each of these subcomponents of a cell also interacts with other sets of macromolecules, creating a large network of protein–protein and protein–nucleic acid interactions throughout the cell. To understand the cell, therefore, one will need to analyze most of these other interactions as well.
Some idea of the complexity of intracellular protein networks can be gained from a particularly well-studied example described in Chapter 16: the many dozens of proteins that interact with the actin cytoskeleton in the yeast Saccharomyces cerevisiae (see Figure 16-15).
The extent of such protein–protein interactions can also be estimated more generally. In particular, large-scale efforts have been undertaken to detect these interactions using the two-hybrid screening method described in Chapter 8 (see Figure 8-51). Thus, for example, this method is being applied to a large set of the 6000 gene products produced by S. cerevisiae. As expected, the majority of the interactions that have been observed are between proteins in the same functional group. That is, proteins involved in cell-cycle control tend to interact extensively with each other, as do proteins involved with DNA synthesis, or DNA repair, and so on. But there are also a surprisingly large number of interactions between the protein members of different functional groups (Figure 3-78). These interactions are presumably important for coordinating cell functions, but most of them are not understood.
A map of the protein–protein interactions observed between different functional groups of proteins in the yeast S. cerevisiae. To produce this map, more than 1500 proteins were assigned to the indicated 31 functional groups. About 70 percent of (more...)
An examination of the range of available data suggests that an average protein in a human cell may interact with somewhere between 5 and 15 different partners. Often, a different set of partners will be bound by each of the different domains in a multidomain protein; in fact, one can speculate that the unusually extensive multidomain structures observed for human proteins (see p. 462) may have evolved to generate these interactions. Given the enormous complexity of the interacting networks of macromolecules in cells, deciphering their full functional meaning may well keep scientists busy for centuries.
Proteins can form enormously sophisticated chemical devices, whose functions largely depend on the detailed chemical properties of their surfaces. Binding sites for ligands are formed as surface cavities in which precisely positioned amino acid side chains are brought together by protein folding. In the same way, normally unreactive amino acid side chains can be activated to make and break covalent bonds. Enzymes are catalytic proteins that greatly speed up reaction rates by binding the high-energy transition states for a specific reaction path; they also perform acid catalysis and base catalysis simultaneously. The rates of enzyme reactions are often so fast that they are limited only by diffusion; rates can be further increased if enzymes that act sequentially on a substrate are joined into a single multienzyme complex, or if the enzymes and their substrates are confined to the same compartment of the cell.
Proteins reversibly change their shape when ligands bind to their surface. The allosteric changes in protein conformation produced by one ligand affect the binding of a second ligand, and this linkage between two ligand-binding sites provides a crucial mechanism for regulating cell processes. Metabolic pathways, for example, are controlled by feedback regulation: some small molecules inhibit and other small molecules activate enzymes early in a pathway. Enzymes controlled in this way generally form symmetric assemblies, allowing cooperative conformational changes to create a steep response to changes in the concentrations of the ligands that regulate them.
Changes in protein shape can be driven in a unidirectional manner by the expenditure of chemical energy. By coupling allosteric shape changes to ATP hydrolysis, for example, proteins can do useful work, such as generating a mechanical force or moving for long distances in a single direction. The three-dimensional structures of proteins, determined by x-ray crystallography, have revealed how a small local change caused by nucleoside triphosphate hydrolysis is amplified to create major changes elsewhere in the protein. By such means, these proteins can serve as input–output devices that transmit information, as assembly factors, as motors, or as membrane-bound pumps. Highly efficient protein machines are formed by incorporating many different protein molecules into larger assemblies in which the allosteric movements of the individual components are coordinated. Such machines are now known to perform many of the most important reactions in cells.
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