Most processes in cells do not take place independently of one another or at a constant rate. The activities of all proteins and other biomolecules are regulated to integrate their functions for optimal performance for survival. For ex ample, the catalytic activity of enzymes is regulated so that the amount of reaction product is just sufficient to meet the needs of the cell. As a result, the steady-state concentrations of substrates and products may vary depending on cellular conditions. Regulation of nonenzymatic proteins—the opening or closing of membrane channels or the assembly of a macromolecular complex, for example—is also essential.
In general, there are three ways to regulate protein activity. First, cells can increase or decrease the steady-state level of the protein by altering its rate of synthesis, its rate of degradation, or both. Second, cells can change the intrinsic activity, as distinct from the amount, of the protein. For example, through noncovalent and covalent interactions, cells can change the affinity of substrate binding, or the fraction of time the protein is in an active versus an inactive conformation. Third, there can be a change in the location or the concentration within the cell of the protein itself, of the target of the protein’s activity (e.g., an enzyme’s substrate), or of some other molecule required for the protein’s activity (e.g., an enzyme’s cofactor). All three types of regulation play essential roles in the lives and functions of cells. In this section, we first discuss mechanisms for regulating the amount of a protein, then turn to noncovalent and covalent interactions that regulate protein activity.
Regulated Synthesis and Degradation of Proteins Is a Fundamental Property of Cells
The rate of synthesis of a protein is determined by the rate at which the DNA encoding the protein is converted to mRNA (transcription), the steady-state amount of the active mRNA in the cell, and the rate at which the mRNA is converted into newly synthesized protein (translation).
The life spans of intracellular proteins vary from as short as a few minutes for mitotic cyclins, which help regulate passage through the mitotic stage of cell division, to as long as the age of an organism for proteins in the lens of the eye. Protein life span is controlled primarily by regulated protein degradation.
Protein degradation plays two especially important roles in the cell. First, it removes proteins that are potentially toxic, improperly folded or assembled, or damaged— including the products of mutated genes and proteins damaged by chemically active cell metabolites or stress (e.g., heat shock). Despite the existence of chaperone-mediated protein folding, some newly made proteins are rapidly degraded because they are misfolded. This degradation might be necessary due to failure of timely engagement of the necessary chaperones to guide the folding of the proteins or due to their defective assembly into complexes. Most other proteins are degraded more slowly, undergoing about 1–2 percent degradation per hour in mammalian cells. Second, the controlled destruction of otherwise normal proteins, along with controlled rates of synthesis, provides a powerful mechanism for maintaining the appropriate levels of the proteins and their activities and for permitting rapid changes in these levels to help the cells respond to changing conditions.
Eukaryotic cells have several pathways for degrading proteins. One major pathway is degradation by enzymes within lysosomes, membrane-limited organelles whose acidic interior (pH ∼4.5) is filled with a host of hydrolytic enzymes. Lysosomal degradation is directed primarily toward aged or defective organelles of the cell—a process called autophagy—and toward extracellular proteins taken up by the cell. Lysosomes will be discussed at length in later chapters. Here we focus on another important degradation pathway: cytoplasmic protein degradation by proteasomes, which can account for up to 90 percent of the protein degradation in mammalian cells.
The Proteasome Is a Molecular Machine Used to Degrade Proteins
Proteasomes are very large protein-degrading molecular machines that influence many different cellular functions, including the cell cycle, transcription, DNA repair, programmed cell death, or apoptosis, recognition of and response to infection by foreign organisms, and removal of misfolded proteins. There are approximately 30,000 proteasomes in a typical mammalian cell.
Proteasomes consist of roughly 60 protein subunits and have a mass of about 2.4 × 106 Da. Proteasomes have a cylindrical, barrel-like catalytic core (Figure 1a), called the 20S proteasome (where S is a Svedberg unit based on the sedimentation properties of the particle and is proportional to its size), which is approximately 14.8 nm tall and 11.3 nm in diameter. Bound to the ends of this core are either one or two 19S cap complexes that regulate the activity of the 20S catalytic core. When the core and one or two caps are combined, they are referred to collectively as the 26S complex, even though the two-cap-containing complex is larger (30S). A 19S cap has 19 protein subunits, six of which can hydrolyze ATP (i.e., they are AAA-type ATPases) to provide the energy needed to unfold protein substrates and selectively transfer them into the inner chamber of the proteasome’s catalytic core. Genetic studies in yeast have shown that cells cannot survive without functional proteasomes, thus demonstrating their importance. Furthermore, proper proteasomal activity is so important that cells will expend as much as 30 percent of the energy needed to synthesize a protein to degrade it in a proteasome.
The 20S proteasomal catalytic core comprises two inner rings of seven β subunits each, with three proteolytic active sites per ring facing toward the ∼1.7-nm-diameter inner chamber formed by those rings, and two outer rings of seven α subunits each, which limit substrate access (see Figure 1a) via an entry channel that can be opened by the 19S cap. Proteasomes can degrade most proteins thoroughly because the three active sites in each β subunit ring can cleave peptide bonds at hydrophobic residues, acidic residues, or basic residues. Polypeptide substrates must enter the chamber via a regulated ∼1.3-nm-diameter aperture at the center of the outer α subunit rings. In the 26S protea some, the opening of the aperture, which is narrow and often allows the entry of only unfolded proteins, is controlled by ATPases in the 19S cap. These ATPases are responsible for unfolding protein substrates and translocating those unfolded polypeptides into the inner chamber of the catalytic core (Figure 1b, bottom right). The short peptide products of proteasomal digestion (2–24 residues long) exit the chamber and are further degraded rapidly by cytosolic peptidases, eventually being converted to individual (“free”) amino acids. One researcher has quipped that a proteasome is a “cellular chamber of doom” in which proteins suffer a “death by a thousand cuts.”
Fig1. Ubiquitin- and proteasome-mediated proteolysis. (a) Right: The 26S proteasome has a cylindrical structure with a 19S cap at one or both ends of the 20S core particle. The nineteen different subunits of the 19S cap (shown in multiple colors) include six AAA ATPase subunits (Rpt1–6, red), which assemble into a heterohexameric ring; two ubiquitin (Ub) receptors (Rpn10 and Rpn13, yellow); and a deubiquitinase enzyme (DUB, Rpn11, green), which forms a heterodimer with its evolutionarily related counterpart Rpn8. Moreover, the 19S cap contains scaffolding and other proteins (tan). The two 19S caps shown are facing in opposite directions relative to the plane of the page. The 20S core consists of four stacked heptameric rings (∼110 Å diameter × 160 Å long), each containing either α (outer rings) or β (inner rings) subunits (blue). Left: Cutaway view of the 20S core, showing the inner chambers. Proteolysis occurs within the central inner chamber of the core formed by the β rings. (b) Proteins are targeted for proteasomal degradation by polyubiquitinylation. Enzyme E1 is activated by the ATP-dependent attachment of a ubiquitin (Ub) molecule (step 1) and then transfers this Ub molecule to a cysteine residue in E2 (step 2 ). Ubiquitin ligase (E3) transfers the bound Ub molecule on E2 to the side-chain –NH2 of a lysine residue in a target protein, forming an isopeptide bond (step 3 ). Additional Ub molecules are added to the Ub-modified target protein via isopeptide bonds to the previously added Ub by repeating steps 1 – 3 , forming a polyubiq uitin chain (step 4 ). The polyubiquitinylated target is recognized by Ub receptors in the proteasome’s 19S cap (step 5 ), and the Ub groups are removed by the deubiquitinase enzyme (step 6 ). In step 7 ATP hydrolysis enables the six protein (hexameric) ATPase subunits (red) to unfold the substrate and transfer the unfolded protein via a pore in the hexamer into the proteolysis chamber in the 20S core (step 7a ), in some cases coordinately with step 6, and the protein is cleaved into short peptide digestion fragments (step 7b ) that are then released (step 7c ). [Part (a) courtesy of Antje Aufderheide and Friedrich Foerster, data from P. Unverdorben et al., 2014, P. Natl. Acad. Sci. USA 111:(15):5544–5549, PDB ID 4cr2.]
Inhibitors of proteasome function have proved to be exceptionally useful in the laboratory and the clinic. Small-molecule proteasome inhibitors, such as MG132, are used to block proteasomal degradation in the lab and to help evaluate the role of the proteasome and, as we shall see below, polyubiquitinylation in a wide variety of processes. Other small-molecule proteasome inhibitors have been used therapeutically. Because of the global importance of proteasome-mediated protein breakdown in cells, continuous, complete inhibition of proteasomes kills cells. However, partial proteasome inhibition for short intervals is widely used as an approach to cancer chemotherapy, especially to treat multiple myeloma, a cancer involving the abnormal proliferation of antibody (immunoglobulin)-producing cells. The myeloma cells produce abnormally high levels of potentially toxic, aberrant immunoglobulin polypeptide chains, which are degraded by proteasomes. Proteasome inhibition in these cancer cells leads to the buildup of toxic, misfolded immunoglobulin polypeptides within the cells, and thus to cell death. In addition, to survive and grow, myeloma cells require the robust activity of a regulatory protein called NF-κB as well as other “pro-survival” and “pro-proliferation” proteins. In turn, NF-κB can function fully and promote survival and proliferation only when its inhibitor, I-κB, is disengaged and degraded by proteasomes. Partial inhibition of proteasomal activity by a small-molecule inhibitor drug results in increased levels of I-κB and, consequently, reduced NF-κB activity (that is, loss of its protective activity). The cancer cells subsequently undergo less proliferation and die by apoptosis. Thus, multiple myeloma cells are more sensitive to proteasome inhibitors than normal cells. Consequently, controlled administration of proteasome inhibitors, at levels that kill the cancer cells but not normal cells, has proved to be an effective therapy for multiple myeloma.
Ubiquitin Marks Cytosolic Proteins for Degradation in Proteasomes
If proteasomes are to rapidly degrade only those proteins that are either defective or scheduled to be removed, they must be able to distinguish between those proteins that need to be degraded and those that don’t. Cells mark proteins that should be degraded by covalently attaching to them a linear chain of multiple copies of a 76-residue polypeptide called ubiquitin (Ub) that is highly conserved from yeast to humans. This “polyubiquitin tail” serves as a cellular “kiss of death,” marking the protein for destruction in the proteasome. The ubiquitinylation process (Figure 3-31b, steps 1–3) involves three distinct steps:
1. Activation of ubiquitin-activating enzyme (E1) by the addition of a ubiquitin molecule, a reaction that requires ATP.
2. Transfer of this ubiquitin molecule to a cysteine residue in a ubiquitin-conjugating enzyme (E2).
3. Formation of a covalent bond between the carboxyl group of the C-terminal glycine 76 of the ubiquitin bound to E2 and the amino group of the side chain of a lysine residue in the target protein, a reaction catalyzed by a ubiquitin-protein ligase (E3). This type of bond is called an isopeptide bond because it covalently links a side-chain amino group, rather than the α amino group, to the carboxyl group. Subsequent ligase reactions covalently attach the C-terminal glycine of an additional ubiquitin molecule via an isopeptide bond to the side chain of lysine 48 of the previously added ubiquitin to generate a polyubiquitin chain covalently attached to the target protein.
Generally, following attachment of four or more ubiquitins in a polyubiquitin chain, the 19S regulatory cap of the 26S proteasome (sometimes with the help of accessory proteins) recognizes the polyubiquitin-labeled protein using its Ub receptors (see Figure 1a), uses ATPases to unfold it, and transports it into the proteasome core for degradation. As a polyubiquitinylated substrate is unfolded and passed into the core of the proteasome, enzymes called deubiquitinases (Dubs) hydrolyze the bonds between the individual ubiquitins and between the targeted protein and ubiquitin, recycling the ubiquitins for additional rounds of protein modification (see Figure1b). Analysis of the human genome sequence indicates the presence of about 90 distinct Dubs, about 80 percent of which use cysteine in a catalytic triad similar to that in the serine proteases described earlier (the sulfhydryl in the cysteine side chain is used in place of the hydroxyl in the side chain of the serine). In some Dubs, zinc is a key participant in the catalytic reactions.
Specificity of Degradation Targeting of specific proteins for proteasomal degradation is primarily achieved through the substrate specificity of E3 ligases (see Figure 1b, step 3). As a testament to their importance, there are an estimated 600 or more ubiquitin ligase genes in the human genome. The many E3 ligases in mammalian cells ensure that the wide variety of proteins to be polyubiquitinylated can be modified when necessary. Some E3 ligases are associated with chaperones that recognize unfolded or misfolded proteins; for example, the E3 ligase CHIP is a co-chaperone for Hsp70. These and other proteins (co-chaperones, escort factors, adapters) can mediate E3 ligase–catalyzed polyubiquitinylation of dysfunctional proteins that cannot be readily refolded properly and, consequently, mediate their delivery to proteasomes for degradation. In such cases, the chaperone-ubiquitinylation-proteasome system works in concert for protein quality control.
In addition to quality control, the ubiquitin-proteasome system can be used to regulate the activity of important cellular proteins. An example is the regulated degradation of proteins called cyclins, which control the cell cycle. Cyclins contain the internal sequence Arg-X X-Leu-Gly-X-Ile-Gly-Asp/Asn (where X can be any amino acid), which is recognized by specific ubiquitinylating enzyme complexes. At a specific time in the cell cycle, each cyclin is phosphorylated by a cyclin kinase. This phosphorylation is thought to cause a conformational change that exposes the recognition sequence to the ubiquitinylating enzymes, leading to polyubiquitinylation and proteasomal degradation.
Other Functions of Ubiquitin and Ubiquitin-Related Molecules There are several close relatives of ubiquitin that employ similar E1-, E2-, and E3-dependent mechanisms of activation and transfer to acceptor substrates. These ubiquitin-like modifiers control processes as diverse as nu clear import, regulated by the ubiquitin-like modifier Sumo, and autophagy, regulated by the ubiquitin-like modifier Atg8/LC3. Furthermore, the attachment of ubiquitin to a target protein can be used for purposes other than to mark the protein for degradation, as we will see later in this section, and some of these functions involve polyubiquitin linkages other than those via Lys-48.
Like ubiquitinylation, deubiquitinylation is involved in processes other than proteasome-mediated protein degradation. Large-scale, mass-spectrometry-based “proteomic” methods described later in this chapter, together with so phisticated computational approaches, have suggested that Dubs, which are often bound in multiprotein complexes, are involved in an extraordinarily wide range of cell processes.
Noncovalent Binding Permits Allosteric, or Cooperative, Regulation of Proteins
In addition to regulating the amount of a protein, cells can also regulate the intrinsic activity of a protein. One of the most important mechanisms for regulating protein function is through allosteric interactions. Broadly speaking, allostery (from the Greek, “other shape”) refers to any change in a protein’s tertiary or quaternary structure, or in both, induced by the noncovalent binding of a ligand. When a ligand binds to one site (A) in a protein and induces a conformational change that alters the activity of a different site (B), the ligand is called an allosteric effector of the protein, while site A is called an allosteric binding site, and the protein is called an allosteric protein. By definition, allosteric proteins have multiple binding sites, at least one for the allosteric effector and at least one for other molecules with which the protein interacts. The allosteric change in activity can be positive or negative; that is, it can be an increase or a decrease in protein activity. Negative allostery often involves the end product of a multi step biochemical pathway binding to, and reducing the activity of, an enzyme that catalyzes an early, rate-controlling step in that pathway. In this way, excessive buildup of the product is prevented. This kind of regulation of a metabolic pathway is also called end-product inhibition or feedback inhibition. Allosteric regulation is particularly prevalent in multimeric enzymes and other proteins in which conformational changes in one subunit are transmitted to an adjacent subunit.
Cooperativity, a term that is often used synonymously with allostery, usually refers to the influence (positive or negative) that the binding of a ligand at one site has on the binding of another molecule of the same type of ligand at a different site. Hemoglobin presents a classic example of positive cooperative binding in that the binding of a single ligand, molecular oxygen (O2), increases the affinity of hemoglobin for the next oxygen molecule. Each of the four subunits in hemoglobin contains one heme molecule. The heme groups are the oxygen-binding components of hemoglobin. The binding of oxygen to the heme molecule in one of the four hemoglobin subunits induces a local conformational change whose effect spreads to the other subunits, lowering the Kd (increasing the affinity) for the binding of additional oxygen molecules to the remaining hemes and yielding a sigmoidal oxygen-binding curve (Figure 2). Because of the sigmoidal shape of the oxygen-saturation curve, it takes only a fourfold increase in oxygen concentration for the saturation of the oxygen-binding sites in hemoglobin to go from 10 to 90 percent. Conversely, if there were no cooperativity and the shape of the curve was typical of that for Michaelis-Men ten, or noncooperative, binding, it would take an eighty-one-fold increase in oxygen concentration to accomplish the same increase in loading of its binding sites in hemoglobin. This cooperativity permits hemoglobin to take up oxygen very efficiently in the lungs, where the oxygen concentration is high, and unload it in tissues, where the con centration is low. Thus cooperativity amplifies the sensitivity of a system to changes in the concentration of its ligands, providing in many cases a selective evolutionary advantage.
Fig2. Hemoglobin binds oxygen cooperatively. Each tetrameric hemoglobin molecule has four oxygen- binding sites; at saturation, all the sites are loaded with oxygen. The oxygen concentration in tissues is commonly measured as the partial pressure of oxygen (pO2) in torr units (a standard measure of pressure equivalent to 1 mm of mercury under standard conditions). P50 is the pO2 at which half the oxygen-binding sites are occupied; it is somewhat analogous to the Km for an enzymatic reaction. The large change in the amount of oxygen bound over a small range of pO2 values permits efficient unloading of oxygen in peripheral tissues such as muscle. The sigmoidal shape of a plot of saturation versus ligand concentration is indicative of cooperative binding, in which the binding of one oxygen molecule allosterically influences the binding of subsequent oxygens. In the absence of cooperative binding, a binding curve is a hyperbola, similar to the curves in Figure 3-24. See J. M. Berg et al., 2015, Biochemistry, 8th ed., Macmillan.
Noncovalent Binding of Calcium and GTP Are Widely Used as Allosteric Switches to Control Protein Activity
Unlike oxygen, which causes graded allosteric changes in the activity of hemoglobin, some other allosteric effectors act as switches, turning the activity of many different proteins on or off by binding to them noncovalently. Two important allosteric switches that we will encounter many times throughout this book, especially in the context of cell signaling pathways, are Ca2+ and GTP.
Ca2+/Calmodulin-Mediated Switching The concentration of Ca2+ that is free in the cytosol (not bound to molecules other than water) is kept very low (∼10−7 M) by specialized membrane transport proteins that continually pump excess Ca2+ out of the cytosol. However, the cytosolic Ca2+ concentration can increase tenfold to a hundredfold when Ca2+-permeable channels in the cell-surface membranes open and allow extracellular Ca2+ to flow into the cell. This rise in cytosolic Ca2+ is sensed by specialized Ca2+-binding proteins, which alter cellular behavior by turning the activities of other proteins on or off. The importance of extracellular Ca2+ for cell activity was first documented by S. Ringer in 1883, when he discovered that isolated rat hearts suspended in an NaCl solution made with “hard” (Ca2+-rich) London tap water contracted beautifully, whereas they beat poorly and stopped quickly if distilled, Ca2+-depleted, water was used.
Many Ca2+-binding proteins bind Ca2+ using the EF hand/helix-loop-helix structural motif discussed earlier. A well-studied EF hand protein, calmodulin, is found in all eukaryotic cells, where it may exist as an individual monomeric protein or as a subunit of a multimeric protein. This dumbbell-shaped molecule contains four Ca2+-binding EF hands with Kds of about 10−6 M. The binding of Ca2+ to calmodulin causes a conformational change that permits Ca2+/calmodulin to bind to conserved sequences in various target proteins (Figure 3), thereby switching their activities on or off. Calmodulin and similar EF hand proteins thus function as switch proteins, acting in concert with changes in Ca2+ levels to modulate the activity of other proteins.
Fig3. Conformational changes induced by Ca2+ binding to calmodulin. Calmodulin is a widely distributed cytosolic protein that contains four Ca2+-binding sites, one in each of its EF hand (helix loop-helix) motifs (EF1-ER4). At cytosolic Ca2+ concentrations above about 5 × 10−7 M, binding of Ca2+ to calmodulin changes the protein’s conformation from the dumbbell-shaped, Ca2+ free form (a) to one in which hydrophobic side chains become more exposed to solvent. The resulting Ca2+/calmodulin complex can wrap around exposed helices (target peptides) with specialized sequences in various target proteins (b), thereby altering their activities. [Part (a) data from H. Kuboniwa et al., 1995, Nat. Struct. Biol. 2:768–776, PDB ID 1cfd. Part (b) data from W. E. Meador, A. R. Means, and F. A. Quiocho, 1992, Science 257:1251–1255, PDB ID 1cdl.]
Switching Mediated by Guanine Nucleotide–Binding Proteins Another group of intracellular switch proteins constitutes the GTPase superfamily. As the name suggests, these proteins are enzymes—GTPases—that can hydrolyze GTP (guanosine triphosphate) to GDP (guanosine diphosphate). They include the monomeric Ras protein with bound GDP and the Gα subunit of the trimeric G proteins. Both Ras and Gα can bind to the plasma membrane, function in cell signaling, and play key roles in cell proliferation and differentiation. Other members of the GTPase superfamily function in protein synthesis, the transport of proteins between the nucleus and the cytoplasm, the formation of coated vesicles and their fusion with target membranes, and rearrangements of the actin cytoskeleton. Some GTPase proteins have a covalently attached lipid chain that mediates their binding to membranes. We examine the roles of various GTPase switch proteins in regulating intracellular signaling and other processes in several later chapters.
All the GTPase switch proteins exist in two forms, or conformations (Figure 4): (1) an active (“on”) form with bound GTP, which can influence the activity of specific target proteins to which they bind, and (2) an inactive (“off”) form with bound GDP. The switch is turned on—that is, the conformation of the protein changes from inactive to active—when a GTP molecule replaces a bound GDP in the inactive conformation. The switch is turned off when the relatively slow GTPase activity of the protein hydrolyzes bound GTP, converting it to GDP and leading the conformation to change to the inactive form. The amount of time any given GTPase switch remains in the active, GTP-bound form depends on the rate of its GTPase activity. Thus the GTPase activity acts as a timer to control this switch. Cells contain a variety of proteins that can modulate the baseline (or intrinsic) rate of GTPase activity for any given GTPase switch and so can control how long the switch remains on.
Fig4. The GTPase switch. GTPases are enzymes that bind to GTP and hydrolyze it to GDP. When bound to GTP, the GTPase protein adopts its active, or “on,” conformation and can interact with target proteins to regulate their activities. When the bound GTP is hydrolyzed to GDP by the intrinsic GTPase activity of the protein, the GTPase with GDP bound assumes an inactive, or “off,” conformation. The GTPase switch can be turned back on when another protein, called a GEF (guanine nucleotide exchange factor), mediates the replacement (exchange) of the bound GDP with a GTP molecule from the surround ing fluid. GTPase-activating proteins, or GAPs, can influence the rates of GTP hydrolysis. The binding of the active form of the GTPase to its targets is a form of noncovalent regulation.
For example, GTPase-activating proteins, or GAPs, increase the rate of GTPase activity, thus reducing the time the GTPase is in the active form. Cells also have specific proteins whose function is to regulate the conversion of inactive GTPases to active ones—that is, to turn the switch on—by mediating the replacement of bound GDP with GTP (GDP/ GTP exchange). These proteins are called guanine nucleotide exchange factors, or GEFs. GTPases with lipid anchors are also regulated by proteins called guanine nucleotide dis sociation inhibitors (GDIs) that bind to the lipid chain and thus influence interactions with cellular membranes.
The GAPs, GEFs, and GDIs are themselves subject to regulation and, together with their GTPases, participate in complex regulatory networks that control a vast array of cellular activities. It is, therefore, not surprising that disruptions of these finely tuned regulatory networks by mutations or pathogens are associated with a wide variety of diseases. Examples of genetic diseases affecting these networks include Noonan syndrome (a developmental disorder), retinitis pigmentosa (a degenerative eye disease), and X-linked mental retardation. Examples of disruptions of these networks by pathogens include bacterially induced food poisoning, dysenteries (inflammation of the intestines with diarrhea), Legionnaires’ disease (a severe type of pneumonia that involves lung inflammation), and even the plague [also called the Black Death, which between 1347 and 1351 decimated the populations of China (∼50 percent death rate) and Europe (∼33 percent death rate)]
Phosphorylation and Dephosphorylation Covalently Regulate Protein Activity
In addition to exploiting the noncovalent regulators described above, cells can use covalent modifications to regulate the intrinsic activity of a protein. One of the most common covalent mechanisms for regulating protein activity is phosphorylation, the reversible addition of phosphate groups to hydroxyl groups on the side chains of serine, threonine, or tyrosine residues. Phosphorylated proteins are called phosphoproteins. Phosphorylation is catalyzed by enzymes called protein kinases, while the removal of phosphates, known as dephosphorylation, is catalyzed by phosphatases. The counteracting activities of kinases and phosphatases provide cells with a “switch” that can turn on or turn off the function of various proteins that are the substrates (or targets) of these enzymes (Figure 5). Sometimes phosphorylation sites are masked transiently by reversible covalent modification with the sugar N-acetylglucosamine (called O-GlcNAcylation), which is an additional means of covalent regulation. Phosphorylation changes a protein’s charge and to some extent its surface shape; it can also result in conformational changes. As a consequence, phosphorylation (or dephosphorylation) can influence the location of a protein within cells (e.g., its attachment to the inner surface of the plasma membrane), its intrinsic (e.g., enzymatic) activity, its ability to bind to other molecules, including metabolites, DNA, or other proteins, its ability to undergo further covalent modification, or its stability (rate of degradation). In addition, several conserved protein domains, such as the SH2 domain, bind specifically to phosphorylated peptides. Thus phosphorylation can mediate the formation of protein complexes that can generate or extinguish a wide variety of cellular activities, discussed in many subsequent chapters.
Fig5. Regulation of protein activity by phosphorylation and dephosphorylation. The cyclic phosphorylation and dephosphorylation of a protein is a common cellular mechanism for regulating protein activity. In this example, the target protein is active (top) when phosphorylated and inactive (bottom) when dephosphorylated; some proteins have the opposite response to phosphorylation.
Nearly 3 percent of all yeast proteins are protein kinases or phosphatases, indicating the importance of phosphorylation and dephosphorylation reactions even in these simple cells. Analysis of the human genome indicates there are approximately 500 human protein kinases (the human “kinome”). All classes of proteins—including structural proteins, scaffolds, enzymes, membrane channels, and signaling molecules—have members regulated by kinase/phosphatase modifications. Different protein kinases and phosphatases are specific for different target proteins, often recognizing different linear sequences in which the residue to be phosphorylated is embedded, and so can regulate distinct cellular pathways, as discussed in later chapters. Some kinases have many targets, so that a single kinase can serve to integrate the activities of many targets simultaneously. Frequently, the target of a kinase or phosphatase is yet another kinase or phosphatase, creating a cascade effect. There are many examples of such kinase cascades, which permit amplification of a signal and many levels of fine-tuning.
Ubiquitinylation and Deubiquitinylation Covalently Regulate Protein Activity
Both ubiquitin and ubiquitin-like proteins (of which there are more than a dozen in humans) can be covalently linked to a target protein in a regulated fashion, in a manner analogous to phosphorylation. Deubiquitinases can reverse ubiquitinylation in a manner analogous to the action of phosphatases. These ubiquitin modifications are structurally far more com plex than phosphorylation, however, and so can mediate many distinct interactions between the ubiquitinylated protein and other cellular proteins. Ubiquitinylation can involve attachment of a single ubiquitin to a protein (monoubiquitinylation), addition of multiple, single ubiquitin molecules to different sites on one target protein (multiubiquitinylation), or addition of a polymeric chain of ubiquitins to a protein (polyubiquitinylation). An additional source of variation is that different amino groups in the ubiquitin molecule can be used to form an isopeptide bond with the C-terminal Gly-76 in another ubiquitin to form a polyubiquitin chain. All seven lysine residues in ubiquitin (Lys-6, Lys-11, Lys-27, Lys-29, Lys-33, Lys-48, and Lys-63) and its N-terminal amino group can participate in inter-ubiquitin linkages. Different ubiquitin ligases are specific both for targets (substrates) to be ubiquitinylated and for the lysine side chains on the ubiquitins that participate in the inter-ubiquitin isopeptide link ages (Lys-63 or Lys-48, etc.) ( Figure 6). These multiple forms of ubiquitinylation result in the generation of a wide variety of recognition surfaces that can participate in many protein-protein interactions with the hundreds of proteins (>200 in humans) that contain more than a dozen distinct ubiquitin-binding domains (UBD). In addition, any given polyubiquitin chain has the potential to bind simultaneously to more than one UBD-containing protein, leading to the formation of ubiquitinylation-dependent multiprotein complexes. Some deubiquitinases can remove an intact polyubiquitin chain from a modified protein (“anchored” chain) and thus generate a polyubiquitin chain not covalently linked to another protein (“unanchored” chain). Even these unanchored chains may serve a regulatory role. With this great structural diversity, it is not surprising that cells use ubiquitinylation and deubiquitinylation to control many different cellular functions.
Fig6. Determination of polyubiquitin function by the lysine used for inter-ubiquitin isopeptide bonds. Different ubiquitin ligases catalyze polyubiquitinylation of distinct target (substrate) proteins (colored ovals) using distinct lysine side chains of ubiquitin molecules (purple) to generate the inter-ubiquitin isopeptide linkages (blue) with Gly-76 of the adjacent ubiquitin. Dotted blue arrows represent additional ubiquitins in the chain that are not shown. The lysine used for the isopeptide bonds determines the function of the polyubiquitinylation. For example, polyubiquitins with Lys-48:Gly-76 isopeptide bonds direct the target to proteasomes for degradation. Those that use Lys-63, Lys-33, and Lys-11 influence signaling, T-lymphocyte control, and cell division, respectively. Isopeptide bonds involving ubiquitin’s Lys-6, Lys-27, and Lys-29 and bonds using its N-terminal amino group (not shown) can also be used to generate polyubiquitin chains.
We have already seen how polyubiquitinylation via Lys-48 residues is used to tag proteins for proteasomal degradation.
There is evidence that polyubiquitinylation via other Lys residues (for example, Lys-11 and Lys-33, but not Lys-63) can also target proteins for proteasomal degradation. Strik ingly, ubiquitinylation unrelated to protein degradation can also control diverse cell functions, including cellular internalization of molecules via endocytosis, repair of damaged DNA, metabolism, messenger RNA synthesis (transcription), defense against pathogens, cell division/cell cycle progression, cell signaling pathways, trafficking of proteins within a cell, and apoptosis. The lysine used to form the inter-ubiquitin isopeptide bonds can vary depending on the cellular system that is regulated (see Figure 6). For example, polyubiquitinylation with Lys-63 linkages is used in many cellular identification and signaling systems, such as recognition of the presence of intracellular viruses and bacteria and the consequent induction of a protective immune response, as well as direction of these pathogens to lysosomes for degradation. Lys-11-linked polyubiquitin chains regulate cell division. Lys-33-linked chains help suppress the activity of receptors on specialized white blood cells, called T lymphocytes, and so control the activity and function of those lymphocytes.
Proteolytic Cleavage Irreversibly Activates or Inactivates Some Proteins
Unlike phosphorylation and ubiquitinylation, which are reversible, the activation or inactivation of protein function by proteolytic cleavage is an irreversible mechanism for regulating protein activity. For example, many polypeptide hormones, such as insulin, are synthesized as longer precursors, and prior to secretion from cells some of their peptide bonds must be hydrolyzed for them to fold properly. In some cases, a single long precursor prohormone polypeptide is cleaved into several distinct active hormones. To prevent the pancreatic serine proteases from inappropriately digesting proteins before they reach the small intestine, they are synthesized as zymogens, inactive precursor enzymes. Cleavage of a peptide bond near the N-terminus of trypsinogen (the zymogen of trypsin) by a highly specific protease in the small intestine generates a new N-terminal residue (Ile-16), whose amino group can form an ionic bond with the carboxylic acid side chain of an internal aspartic acid. This binding causes a con formational change that opens the substrate-binding site, activating the enzyme. The active trypsin can then activate trypsinogen, chymotrypsinogen, and other zymogens. Simi lar but more elaborate protease cascades (with one protease activating inactive precursors of others) that can amplify an initial signal play important roles in several systems, such as the blood-clotting cascade and the complement system. The importance of carefully regulating such systems is clear—inappropriate clotting, for example, could fatally clog the circulatory system, while insufficient clotting could lead to uncontrolled bleeding.
An unusual and rare type of proteolytic processing, termed protein self-splicing, takes place in bacteria and some eukaryotes. This process is analogous to editing film: an internal segment of a polypeptide is removed and the ends of the polypeptide are rejoined (ligated). Unlike other forms of proteolytic processing, protein self-splicing is an autocatalytic process, which proceeds by itself without the participation of other enzymes. The excised peptide appears to eliminate itself from the protein by a mechanism simi lar to that used in the processing of some RNA molecules. In vertebrate cells, the processing of some proteins includes self-cleavage, but the subsequent ligation step is absent. One such protein is Hedgehog, a membrane bound signaling molecule that is critical to a number of developmental processes.
Higher-Order Regulation Includes Control of Protein Location
All the regulatory mechanisms heretofore described affect a protein locally at its site of action, altering the protein’s con centration or turning its activity on or off. Normal functioning of a cell, however, also requires the segregation of proteins to particular compartments, such as the mitochondria, nucleus, or lysosomes. In regard to enzymes, compartmentation not only provides an opportunity for controlling the delivery of substrate or the exit of product, but also permits competing reactions to take place simultaneously in different parts of a cell.
