第五章 蛋白质的性质、分离纯化和鉴定

The amino and carboxyl groups of amino acids, along with the ionizable R groups of some amino acids, function as weak acids and bases. When an amino acid lacking an ionizable R group is dissolved in water at neutral pH, the α-amino and carboxyl groups create a dipolar ion, or zwitterion (German for “hybrid ion”), which can act as either an acid or a base (Fig. 3-9). Substances having this dual (acid-base) nature are amphoteric and are often called ampholytes (from “amphoteric electrolytes”).

Peptides contain only one free α-amino group and one free α-carboxyl group, at opposite ends of the chain (Fig. 3-15). These groups ionize as they do in free amino acids. The α-amino and α-carboxyl groups of all nonterminal amino acids are covalently joined in the peptide bonds. They can no longer ionize and thus do not contribute to the total acid-base behavior of peptides. Ionizable R groups in a peptide (Table 3-1) also contribute to the overall acid-base properties of the molecule (Fig. 3-15).

Like free amino acids, peptides have characteristic titration curves and a characteristic isoelectric pH (pI) at which the net charge is zero and they do not move in an electric field.

As is evident in Figure 3-10, glycine has a net negative charge at any pH above its pI and thus will move toward the positive electrode (the anode) when placed in an electric field. At any pH below its pI, glycine has a net positive charge and will move toward the negative electrode (the cathode). The farther the pH of a glycine solution is from its isoelectric point, the greater the net electric charge of the population of glycine molecules.

Finally, in an aqueous environment, only histidine has an R group (pKa = 6.0) providing significant buffering power near the neutral pH usually found in the intracellular and extracellular fluids of most animals and bacteria (Table 3-1).

Water is a polar solvent. It readily dissolves most biomolecules, which are generally charged or polar compounds (Table 2-1); compounds that dissolve easily in water are hydrophilic (from the Greek, meaning “water-loving”). In contrast, nonpolar solvents such as chloroform and benzene are poor solvents for polar biomolecules but easily dissolve those that are hydrophobic—nonpolar molecules such as lipids and waxes.

Water dissolves salts such as NaCl by hydrating and stabilizing the Na+ and Cl− ions, weakening the electrostatic interactions between them and thus counteracting their tendency to associate in a crystalline lattice (Fig. 2-6).

On carefully examining the contribution of weak interactions to protein stability, we find that the hydrophobic effect generally predominates. Pure water contains a network of hydrogen-bonded H2O molecules. No other molecule has the hydrogen-bonding potential of water, and the presence of other molecules in an aqueous solution disrupts the hydrogen bonding of water. When water surrounds a hydrophobic molecule, the optimal arrangement of hydrogen bonds results in a highly structured shell, or solvation layer, of water around the molecule (see Fig. 2-7). The increased order of the water molecules in the solvation layer correlates with an unfavorable decrease in the entropy of the water.

Water molecules tend to move from a region of higher water concentration to one of lower water concentration, in accordance with the tendency in nature for a system to become disordered. When two different aqueous solutions are separated by a semipermeable membrane (one that allows the passage of water but not solute molecules), water molecules diffusing from the region of higher water concentration to the region of lower water concentration produce osmotic pressure (Fig. 2-11).

Proteins are separated and purified on the basis of differences in their properties. Proteins can be selectively precipitated by changes in pH or temperature, and particularly by the addition of certain salts. A wide range of chromatographic procedures makes use of differences in size, binding affinities, charge, and other properties.

Ionic interactions between dissolved ions are much stronger in less polar environments, because there is less screening of charges by the nonpolar solvent. Water is effective in screening the electrostatic interactions between dissolved ions because it has a high dielectric constant, a physical property that reflects the number of dipoles in a solvent.

For water at 25 °C, ε is 78.5, and for the very nonpolar solvent benzene, ε is 4.6. The dependence on r2 is such that ionic attractions or repulsions operate only over short distances—in the range of 10 to 40 nm (depending on the electrolyte concentration) when the solvent is water.

Denaturation often leads to protein precipitation, a consequence of protein aggregate formation as exposed hydrophobic surfaces associate. The aggregates are often highly disordered. One example is the protein precipitate that can be seen after an egg white is boiled. More-ordered aggregates are also observed in some proteins, as we shall see.

Protein structures have evolved to function in particular cellular or extracellular environments. Conditions different from these environments can result in protein structural changes, large and small. A loss of three-dimensional structure sufficient to cause loss of function is called denaturation. The denatured state does not necessarily equate with complete unfolding of the protein and randomization of conformation. Under most conditions, denatured proteins exist in a set of partially folded states.

Stability is the tendency of a protein to maintain a native conformation. Native proteins are only marginally stable; the ΔG separating the folded and unfolded states in typical proteins under physiological conditions is in the range of only 5 to 65 kJ/mol. A given polypeptide chain can theoretically assume countless conformations, and as a result, the unfolded state of a protein is characterized by a high degree of conformational entropy. This entropy, along with the hydrogen-bonding interactions of many groups in the polypeptide chain with the solvent (water), tends to maintain the unfolded state.

The abruptness of the change suggests that unfolding is a cooperative process: loss of structure in one part of the protein destabilizes other parts. The effects of heat on proteins can be mitigated by structure. The very heat-stable proteins of thermophilic bacteria and archaea have evolved to function at the temperature of hot springs (∼100 °C). The folded structures of these proteins are often similar to those of proteins in other organisms, but take some of the principles outlined here to extremes. They often feature high densities of charged residues on their surfaces, even tighter hydrophobic packing in their interiors, and folds rendered less flexible by networks of ion pairs.

Proteins can also be denatured by extremes of pH, by certain miscible organic solvents such as alcohol or acetone, by certain solutes such as urea and guanidine hydrochloride, or by detergents. Each of these denaturing agents represents a relatively mild treatment in the sense that no covalent bonds in the polypeptide chain are broken. Organic solvents, urea, and detergents act primarily by disrupting the hydrophobic aggregation of nonpolar amino acid side chains that produces the stable core of globular proteins; urea also disrupts hydrogen bonds; and extremes of pH alter the net charge on a protein, causing electrostatic repulsion and the disruption of some hydrogen bonding. The denatured structures resulting from these various treatments are not necessarily the same.

The tertiary structure of a globular protein is determined by its amino acid sequence. The most important proof of this came from experiments showing that denaturation of some proteins is reversible. Certain globular proteins denatured by heat, extremes of pH, or denaturing reagents will regain their native structure and their biological activity if they are returned to conditions in which the native conformation is stable. This process is called renaturation.

A classic example is the denaturation and renaturation of ribonuclease A, demonstrated by Christian Anfinsen in the 1950s. Purified ribonuclease A denatures completely in a concentrated urea solution in the presence of a reducing agent. The reducing agent cleaves the four disulfide bonds to yield eight Cys residues, and the urea disrupts the stabilizing hydrophobic effect, thus freeing the entire polypeptide from its folded conformation. Denaturation of ribonuclease is accompanied by a complete loss of catalytic activity. When the urea and the reducing agent are removed, the denatured ribonuclease spontaneously refolds into its correct tertiary structure, with full restoration of its catalytic activity (Fig. 4-25). The refolding of ribonuclease is so accurate that the four intrachain disulfide bonds are re-formed in the same positions in the renatured molecule as in the native ribonuclease.

The Anfinsen experiment provided the first evidence that the amino acid sequence of a polypeptide chain contains all the information required to fold the chain into its native, threedimensional structure. Subsequent work has shown that only a minority of proteins, many of them small and inherently stable, will fold spontaneously into their native form. Even though all proteins have the potential to fold into their native structure, many require some assistance.

A pure preparation is usually essential before a protein’s properties and activities can be determined. Given that cells contain thousands of different kinds of proteins, how can one protein be purified?

The source of a protein is generally tissue or microbial cells. The first step in any protein purification procedure is to break open these cells, releasing their proteins into a solution called a crude extract. If necessary, differential centrifugation can be used to prepare subcellular fractions or to isolate specific organelles (see Fig. 1-7).

The advent of genetic engineering approaches has provided new and simpler paths for protein purification. The latter methods, described in Chapter 9, often artificially modify the protein being purified, adding a few or many amino acid residues to one or both ends. In many cases, the modifications alter protein function. Isolation of unaltered native proteins requires removal of the modification or a reliance on methods described here.

Commonly, the extract is subjected to treatments that separate the proteins into different fractions based on a property such as size or charge; the process is referred to as fractionation. Early fractionation steps in a purification utilize differences in protein solubility, which is a complex function of pH, temperature, salt concentration, and other factors. The solubility of proteins is lowered in the presence of some salts, an effect called “salting out.” Ammonium sulfate ((NH4)2SO4) is particularly effective for selectively precipitating some proteins while leaving others in solution.

As a salt such as NaCl dissolves, the Na+ and Cl− ions leaving the crystal lattice acquire far greater freedom of motion (Fig. 2-6). The resulting increase in entropy (randomness) of the system is largely responsible for the ease of dissolving salts such as NaCl in water. In thermodynamic terms, formation of the solution occurs with a favorable free-energy change: ΔG = ΔH − TΔS, where ΔH has a small positive value and TΔS a large positive value; thus ΔG is negative.

A solution containing the protein of interest usually must be further altered before subsequent purification steps are possible. For example, dialysis is a procedure that separates proteins from small solutes by taking advantage of the proteins’ larger size.

The membrane allows the exchange of salt and buffer but not proteins. Thus dialysis retains large proteins within the membranous bag or tube while allowing the concentration of other solutes in the protein preparation to change until they come into equilibrium with the solution outside the membrane. Dialysis might be used, for example, to remove ammonium sulfate from the protein preparation.

The choice of methods is somewhat empirical, and many strategies may be tried before the most effective one is found. Researchers can often minimize trial and error by basing the new procedure on purification techniques developed for similar proteins. Common sense dictates that inexpensive procedures such as salting out be used first, when the total volume and the number of contaminants are greatest. As each purification step is completed, the sample size generally becomes smaller (Table 3-5), making it feasible to use more sophisticated (and expensive) chromatographic procedures at later stages.

A porous solid material with appropriate chemical properties (the stationary phase) is held in a column, and a buffered solution (the mobile phase) migrates through it. The protein, dissolved in the same buffered solution that was used to establish the mobile phase, is layered on the top of the column. The protein then percolates through the solid matrix as an everexpanding band within the larger mobile phase. Individual proteins migrate faster or more slowly through the column, depending on their properties.

Protein purification protocols often use genetic engineering to fuse additional amino acids or peptides (tags) to the target protein. Affinity chromatography can be used to bind this tag, achieving a large increase in purity in a single step (see Fig. 9-11). In many cases, the tag can be subsequently removed, fully restoring the function of the native protein.

Enzymes are usually assayed at their optimum pH and at some convenient temperature within the range 25 to 38 ∘C. Also, very high substrate concentrations are generally used so that the initial reaction rate, measured experimentally, is proportional to enzyme concentration (Chapter 6).

By international agreement, 1.0 unit of enzyme activity for most enzymes is defined as the amount of enzyme causing transformation of 1.0 μmol of substrate to product per minute at 25 °C under optimal conditions of measurement (for many enzymes, this definition is inconvenient, and a unit may be defined differently). The term activity refers to the total units of enzyme in a solution. The specific activity is the number of enzyme units per milligram of total protein (Fig. 3-22). The specific activity is a measure of enzyme purity: it increases during purification of an enzyme and becomes maximal and constant when the enzyme is pure (Table 3-5).

After each purification step, the activity of the preparation (in units of enzyme activity) is assayed, the total amount of protein is determined independently, and the ratio of the two gives the specific activity. Activity and total protein generally decrease with each step. Activity decreases because there is always some loss due to inactivation or nonideal interactions with chromatographic materials or other molecules in the solution. Total protein decreases because the objective is to remove as much unwanted or nonspecific protein as possible.

A purification table documents the success of each step in a purification protocol. In the hypothetical purification shown in Table 3-5, the ratio of the final specific activity (15,000 units/mg) to the starting specific activity (10 units/mg) gives the purification factor (1,500). The percentage of the total activity at the last step (45,000 units) relative to the total activity in the starting material (100,000 units) gives the yield from the purification procedure (45%).

The data are assembled in a purification table similar to Table 3-5. A protein is generally considered pure when further purification steps fail to increase specific activity and when only a single protein species can be detected (for example, by electrophoresis in the presence of SDS).

For proteins that are not enzymes, other quantification methods are required. Transport proteins can be assayed by their binding to the molecule they transport, and hormones and toxins by the biological effect they produce; for example, growth hormones will stimulate the growth of certain cultured cells. Some structural proteins represent such a large fraction of a tissue mass that they can be readily extracted and purified without a functional assay. The approaches are as varied as the proteins themselves.

Proteins lead a surprisingly precarious existence. As we have seen, a native protein conformation is only marginally stable. In addition, most proteins must maintain conformational flexibility in order to function.

The noncovalent interactions we have described—hydrogen bonds and ionic, hydrophobic, and van der Waals interactions (Table 2-4)—are much weaker than covalent bonds. An input of about 350 kJ of energy is required to break a mole of (6 × 1023) C—C single bonds, and about 410 kJ is needed to break a mole of C—H bonds, but as little as 4 kJ is sufficient to disrupt a mole of typical van der Waals interactions.

The marginal stability of most proteins can produce a tenuous balance between folded and unfolded states. As proteins are synthesized on ribosomes (Chapter 27), they must fold into their native conformations. Sometimes this occurs spontaneously, but often it requires the assistance of specialized enzymes and complexes called chaperones, which we discuss later in the chapter. Many of these same folding helpers function to refold proteins that become transiently unfolded. Proteins that are not properly folded often have exposed hydrophobic surfaces that render them “sticky,” leading to the formation of inactive aggregates.

Outside the cell, the environment is often more oxidizing, and disulfide formation is more likely to occur. In eukaryotes, disulfide bonds are found primarily in secreted, extracellular proteins (for example, the hormone insulin). Disulfide bonds are also uncommon in bacterial proteins. However, thermophilic bacteria, as well as the archaea, typically have many proteins with stabilizing disulfide bonds; this is presumably an adaptation to life at high temperatures.

Commonly, the extract is subjected to treatments that separate the proteins into different fractions based on a property such as size or charge; the process is referred to as fractionation. Early fractionation steps in a purification utilize differences in protein solubility, which is a complex function of pH, temperature, salt concentration, and other factors.

The advent of genetic engineering approaches has provided new and simpler paths for protein purification. The latter methods, described in Chapter 9, often artificially modify the protein being purified, adding a few or many amino acid residues to one or both ends. In many cases, the modifications alter protein function. Isolation of unaltered native proteins requires removal of the modification or a reliance on methods described here.

The choice of methods is somewhat empirical, and many strategies may be tried before the most effective one is found. Researchers can often minimize trial and error by basing the new procedure on purification techniques developed for similar proteins. Common sense dictates that inexpensive procedures such as salting out be used first, when the total volume and the number of contaminants are greatest. As each purification step is completed, the sample size generally becomes smaller (Table 3-5), making it feasible to use more sophisticated (and expensive) chromatographic procedures at later stages.

Proteins are separated and purified on the basis of differences in their properties. Proteins can be selectively precipitated by changes in pH or temperature, and particularly by the addition of certain salts. A wide range of chromatographic procedures makes use of differences in size, binding affinities, charge, and other properties. These include ion-exchange, size-exclusion, affinity, and high-performance liquid chromatography.

A final important piece of information derived from the titration curve of an amino acid is the relationship between its net charge and the pH of the solution. At pH 5.97, the point of inflection between the two stages in its titration curve, glycine is present predominantly as its dipolar form, fully ionized but with no net electric charge (Fig. 3-10). The characteristic pH at which the net electric charge is zero is called the isoelectric point or isoelectric pH, designated pI.

As is evident in Figure 3-10, glycine has a net negative charge at any pH above its pI and thus will move toward the positive electrode (the anode) when placed in an electric field. At any pH below its pI, glycine has a net positive charge and will move toward the negative electrode (the cathode). The farther the pH of a glycine solution is from its isoelectric point, the greater the net electric charge of the population of glycine molecules.

One method for separating polypeptides makes use of their different solubilities. The solubility of large polypeptides in water depends on the relative polarity of their R groups, particularly on the number of ionized groups: the more ionized groups there are, the more soluble the polypeptides are.

The solubility of proteins is lowered in the presence of some salts, an effect called “salting out.” Ammonium sulfate ((NH4)2SO4) is particularly effective for selectively precipitating some proteins while leaving others in solution. Low-speed centrifugation is then used to remove the precipitated proteins from those remaining in solution.

Water dissolves salts such as NaCl by hydrating and stabilizing the Na+ and Cl− ions, weakening the electrostatic interactions between them and thus counteracting their tendency to associate in a crystalline lattice (Fig. 2-6).

In addition to requiring this input of energy, dissolving hydrophobic compounds in water produces a measurable decrease in entropy. Water molecules in the immediate vicinity of a nonpolar solute are constrained in their possible orientations, as they form a highly ordered cagelike shell around each solute molecule to maximize solvent-solvent hydrogen bonding.

By clustering together, nonpolar regions of the molecules achieve the greatest thermodynamic stability by minimizing the number of ordered water molecules required to surround hydrophobic portions of the solute molecules, increasing the entropy of the system.

The noncovalent interactions we have described—hydrogen bonds and ionic, hydrophobic, and van der Waals interactions (Table 2-4)—are much weaker than covalent bonds. An input of about 350 kJ of energy is required to break a mole of (6 × 1023) C—C single bonds, and about 410 kJ is needed to break a mole of C—H bonds, but as little as 4 kJ is sufficient to disrupt a mole of typical van der Waals interactions. Interactions driven by the hydrophobic effect are also much weaker than covalent bonds, although they are substantially strengthened by a highly polar solvent (a concentrated salt solution, for example).

Water is a polar solvent. It readily dissolves most biomolecules, which are generally charged or polar compounds (Table 2-1); compounds that dissolve easily in water are hydrophilic (from the Greek, meaning “water-loving”). In contrast, nonpolar solvents such as chloroform and benzene are poor solvents for polar biomolecules but easily dissolve those that are hydrophobic—nonpolar molecules such as lipids and waxes.

Ionic interactions between dissolved ions are much stronger in less polar environments, because there is less screening of charges by the nonpolar solvent. Water is effective in screening the electrostatic interactions between dissolved ions because it has a high dielectric constant, a physical property that reflects the number of dipoles in a solvent. The strength, or force (F), of ionic interactions in a solution depends on the magnitude of the charges (Q), the distance between the charged groups (r), and the dielectric constant (ε, which is dimensionless) of the solvent in which the interactions occur:

For water at 25 °C, ε is 78.5, and for the very nonpolar solvent benzene, ε is 4.6. The dependence on r2 is such that ionic attractions or repulsions operate only over short distances—in the range of 10 to 40 nm (depending on the electrolyte concentration) when the solvent is water. In biomolecules, it is not the dielectric constant for the bulk solvent, but the highly localized dielectric constant, as in a hydrophobic pocket of a protein, that determines the interaction of two polar moieties.

A solution containing the protein of interest usually must be further altered before subsequent purification steps are possible. For example, dialysis is a procedure that separates proteins from small solutes by taking advantage of the proteins’ larger size.

The membrane allows the exchange of salt and buffer but not proteins. Thus dialysis retains large proteins within the membranous bag or tube while allowing the concentration of other solutes in the protein preparation to change until they come into equilibrium with the solution outside the membrane. Dialysis might be used, for example, to remove ammonium sulfate from the protein preparation.

The source of a protein is generally tissue or microbial cells. The first step in any protein purification procedure is to break open these cells, releasing their proteins into a solution called a crude extract. If necessary, differential centrifugation can be used to prepare subcellular fractions or to isolate specific organelles (see Fig. 1-7).

The most efficient methods for fractionating proteins make use of column chromatography, which takes advantage of differences in protein charge, size, binding affinity, and other properties (Fig. 3-16). A porous solid material with appropriate chemical properties (the stationary phase) is held in a column, and a buffered solution (the mobile phase) migrates through it. The protein, dissolved in the same buffered solution that was used to establish the mobile phase, is layered on the top of the column. The protein then percolates through the solid matrix as an everexpanding band within the larger mobile phase. Individual proteins migrate faster or more slowly through the column, depending on their properties.

The overall protein band thus widens as it moves through the column. Individual types of proteins (such as A, B, and C, shown in blue, red, and green) gradually separate from each other, forming bands within the broader protein band. Separation improves (i.e., resolution increases) as the length of the column increases. However, each individual protein band also broadens with time due to diffusional spreading, a process that decreases resolution. In this example, protein A is well separated from B and C, but diffusional spreading prevents complete separation of B and C under these conditions. Protein C is being detected and its presence recorded as it is eluted from the column.

As the protein-containing solution exits a column, successive portions (fractions) of this effluent are collected in test tubes. Each fraction can be tested for the presence of the protein of interest as well as other properties, such as ionic strength or total protein concentration. All fractions positive for the protein of interest can be combined as the product of this chromatographic step of the protein purification.

Ion-exchange chromatography exploits differences in the sign and magnitude of the net electric charge of proteins at a given pH (Fig. 3-17a). The column matrix is a synthetic polymer (resin) containing bound charged groups; those with bound anionic groups are called cation exchangers, and those with bound cationic groups are called anion exchangers. The affinity of each protein for the charged groups on the column is affected by the pH (which determines the ionization state of the molecule) and the concentration of competing free salt ions in the surrounding solution. Separation can be optimized by gradually changing the pH and/or salt concentration of the mobile phase in order to create a pH or salt gradient. In cation-exchange chromatography, proteins with a net positive charge migrate through the matrix more slowly than those with a net negative charge, because the migration of the former is retarded more by interaction with the stationary phase.

A biochemist wants to separate two peptides by ion-exchange chromatography. At the pH of the mobile phase to be used on the column, one peptide (A) has a pI of 5.1, due to the presence of more Glu and Asp residues than Arg, Lys, and His residues, and has a net negative charge at neutral pH. Peptide B has a pI of 7.8, reflecting a plurality of positively charged amino acid residues at neutral pH. At neutral pH, which peptide would elute first from a cation-exchange resin? Which would elute first from an anionexchange resin?

A cation-exchange resin has negative charges and binds positively charged molecules, retarding their progress through the column.

Peptide B, with its higher pI and net positive charge, will interact more strongly than peptide A with the cation-exchange resin. Thus, peptide A will elute first. On the anion-exchange resin, peptide B will elute first. Peptide A, having a relatively low pI and a net negative charge, will be retarded by its interaction with the positively charged resin.

Figure 3-17 shows two variations of column chromatography in addition to ion exchange. Size-exclusion chromatography, also called gel filtration (Fig. 3-17b), separates proteins according to size. In this method, large proteins emerge from the column sooner than small ones — a somewhat counterintuitive result. The solid phase consists of cross-linked polymer beads with engineered pores or cavities of a particular size. Large proteins cannot enter the cavities and so take a shorter (and more rapid) path through the column, around the beads. Small proteins enter the cavities and are slowed by their more labyrinthine path through the column. Size-exclusion chromatography can also be used to approximate the size of a protein being purified, using methods similar to those described in Figure 3-19.

(In similar fashion, a set of standard proteins with reproducible retention times on a size-exclusion column can be used to create a standard curve of retention time versus log Mr. The retention time of an unknown substance on the column can be compared with this standard curve to obtain an approximate Mr.)

A protein has a molecular mass of 400 kDa when measured by size-exclusion chromatography. When subjected to gel electrophoresis in the presence of sodium dodecyl sulfate (SDS), the protein gives three bands with molecular masses of 180, 160, and 60 kDa. When electrophoresis is carried out in the presence of SDS and dithiothreitol, three bands again form, this time with molecular masses of 160, 90, and 60 kDa. How many subunits does the protein have, and what is the molecular mass of each?

Affinity chromatography is based on binding affinity (Fig. 3-17c). The beads in the column have a covalently attached chemical group called a ligand — a group or molecule that binds to a macromolecule such as a protein. When a protein mixture is added to the column, any protein with affinity for this ligand binds to the beads, and its migration through the matrix is retarded.

Proteins that do not bind to ATP flow more rapidly through the column. Bound proteins are then eluted by a solution containing either a high concentration of salt or a free ligand — in this case, ATP or an analog of ATP. Salt weakens the binding of the protein to the immobilized ligand, interfering with ionic interactions. Free ligand competes with the ligand attached to the beads, releasing the protein from the matrix; the protein product that elutes from the column is often bound to the ligand used to elute it.

Protein purification protocols often use genetic engineering to fuse additional amino acids or peptides (tags) to the target protein. Affinity chromatography can be used to bind this tag, achieving a large increase in purity in a single step (see Fig. 9-11). In many cases, the tag can be subsequently removed, fully restoring the function of the native protein.

As the crude extract percolates through this matrix, the fusion protein becomes immobilized by binding the glutathione. The other proteins in the extract are washed through the column and discarded. The interaction between GST and glutathione is tight but noncovalent, allowing the fusion protein to be gently eluted from the column with a solution containing either a higher concentration of salts or free glutathione to compete with the immobilized ligand for GST binding. The fusion protein is often obtained with good yield and high purity. In some commercially available systems, the tag can be entirely or largely removed from the purified fusion protein by a protease that cleaves a sequence near the junction between the target protein and its tag.

A shorter tag with widespread application consists of a simple sequence of six or more His residues. These histidine tags, or His tags, bind tightly and specifically to nickel ions. A chromatography matrix with immobilized Ni2+ can be used to quickly separate a His-tagged protein from other proteins in an extract.

Affinity chromatography using terminal tags is powerful and convenient. The tags have been successfully used in thousands of published studies; in many cases, the protein would be impossible to purify and study without the tag. However, even very small tags can affect the properties of the proteins they are attached to, thereby influencing the study results. For example, the tag may adversely affect protein folding. Even if the tag is removed by a protease, one or a few extra amino acid residues can remain behind on the target protein, which may or may not affect the protein’s activity.

When an amphipathic compound (Table 2-1) is mixed with water, the polar, hydrophilic region interacts favorably with the water and tends to dissolve, but the nonpolar, hydrophobic region tends to avoid contact with the water (Fig. 2-7a). The nonpolar regions of the molecules cluster together to present the smallest hydrophobic area to the aqueous solvent, and the polar regions are arranged to maximize their interaction with each other and with the solvent (Fig. 2-7b), a phenomenon called the hydrophobic effect.

Many biomolecules are amphipathic; proteins, pigments, certain vitamins, and the sterols and phospholipids of membranes all have both polar and nonpolar surface regions. Structures composed of these molecules are stabilized by the hydrophobic effect, which favors aggregation of the nonpolar regions. The hydrophobic effect on interactions among lipids, and between lipids and proteins, is the most important determinant of structure in biological membranes. The aggregation of nonpolar amino acids in protein interiors, driven by the hydrophobic effect, also stabilizes the three-dimensional structures of proteins.

Chromatographic methods are typically enhanced by the use of HPLC, or high-performance liquid chromatography. HPLC makes use of high-pressure pumps that speed the movement of the protein molecules down the column; it also uses higherquality chromatographic materials that can withstand the crushing force of the pressurized flow. By reducing the transit time on the column, HPLC can limit diffusional spreading of protein bands and thus can greatly improve resolution.

Protein purification is usually complemented by electrophoresis, an analytical process that allows researchers to visualize and characterize proteins as they are purified. This method does not itself contribute to purification, as electrophoresis often adversely affects the structure and thus the function of proteins. However, it allows a biochemist to rapidly estimate the number of different proteins in a mixture and the degree of purity of a particular protein preparation. Also, electrophoresis can be used to determine such crucial properties of a protein as its isoelectric point and approximate molecular weight.

Like free amino acids, peptides have characteristic titration curves and a characteristic isoelectric pH (pI) at which the net charge is zero and they do not move in an electric field.

Migration may also be affected by protein shape. In electrophoresis, the force moving the macromolecule is the electrical potential, E. The electrophoretic mobility, μ, of a molecule is the ratio of its velocity, V, to the electrical potential. Electrophoretic mobility is also equal to the net charge, Z, of the molecule divided by the frictional coefficient, f, which reflects in part a protein’s shape.

FIGURE 3-18 Electrophoresis. (a) Different samples are loaded in wells or depressions at the top of the SDS polyacrylamide gel. The proteins move into the gel when an electric field is applied. The gel minimizes convection currents caused by small temperature gradients, as well as protein movements other than those induced by the electric field.

The electrophoretic method commonly employed for estimation of purity and molecular weight makes use of the detergent sodium dodecyl sulfate (SDS) (“dodecyl” denoting a 12-carbon chain).

In addition, SDS binding partially unfolds proteins, such that most SDS-bound proteins assume a similar rodlike shape. Electrophoresis in the presence of SDS therefore separates proteins almost exclusively on the basis of mass (molecular weight), with smaller polypeptides migrating more rapidly. After electrophoresis, the proteins are visualized by adding a dye such as Coomassie blue, which binds to proteins but not to the gel itself (Fig. 3-18b). Thus, a researcher can monitor the progress of a protein purification procedure as the number of protein bands visible on the gel decreases after each new fractionation step. When compared with the positions to which proteins of known molecular weight migrate in the gel, the position of an unidentified protein can provide a good approximation of its molecular weight (Fig. 3-19). If the protein has two or more different subunits, generally the subunits are separated by the SDS treatment, and a separate band appears for each.

(b) Proteins can be visualized after electrophoresis by treating the gel with a stain such as Coomassie blue, which binds to the proteins but not to the gel itself. Each band on the gel represents a different protein (or protein subunit); smaller proteins move through the gel more rapidly than larger proteins and therefore are found nearer the bottom of the gel.

Isoelectric focusing is a procedure used to determine the isoelectric point (pI) of a protein (Fig. 3-20).

When a protein mixture is applied, each protein migrates until it reaches the pH that matches its pI. Proteins with different isoelectric points are thus distributed differently throughout the gel.

The amino and carboxyl groups of amino acids, along with the ionizable R groups of some amino acids, function as weak acids and bases. When an amino acid lacking an ionizable R group is dissolved in water at neutral pH, the α-amino and carboxyl groups create a dipolar ion, or zwitterion (German for “hybrid ion”), which can act as either an acid or a base (Fig. 3-9). Substances having this dual (acid-base) nature are amphoteric and are often called ampholytes (from “amphoteric electrolytes”).

Combining isoelectric focusing and SDS electrophoresis sequentially in a process called two-dimensional electrophoresis permits the resolution of complex mixtures of proteins (Fig. 321). This is a more sensitive analytical method than either electrophoretic method alone.

FIGURE 3-21 Two-dimensional electrophoresis. Proteins are first separated by isoelectric focusing in a thin strip gel. The gel is then laid horizontally on a second, slab-shaped gel, and the proteins are separated by SDS polyacrylamide gel electrophoresis. Horizontal separation reflects differences in pI; vertical separation reflects differences in molecular weight. The original protein complement is thus spread in two dimensions. Thousands of cellular proteins can be resolved using this technique. Individual protein spots can be cut out of the gel and identified by mass spectrometry (see Figs 3-28 and 3-29).

Electrophoresis separates proteins on the basis of mass or charge for analytical purposes. SDS gel electrophoresis and isoelectric focusing can be used separately or in combination for higher resolution. All purification procedures require a method for quantifying or assaying the protein of interest in the presence of other proteins. Purification can be monitored by assaying specific activity.

The electrophoretic mobility, μ, of a molecule is the ratio of its velocity, V, to the electrical potential. Electrophoretic mobility is also equal to the net charge, Z, of the molecule divided by the frictional coefficient, f, which reflects in part a protein’s shape.

The electrophoretic method commonly employed for estimation of purity and molecular weight makes use of the detergent sodium dodecyl sulfate (SDS) (“dodecyl” denoting a 12-carbon chain). A protein will bind about 1.4 times its weight of SDS, nearly one molecule of SDS for each amino acid residue.

In addition, SDS binding partially unfolds proteins, such that most SDS-bound proteins assume a similar rodlike shape. Electrophoresis in the presence of SDS therefore separates proteins almost exclusively on the basis of mass (molecular weight), with smaller polypeptides migrating more rapidly. After electrophoresis, the proteins are visualized by adding a dye such as Coomassie blue, which binds to proteins but not to the gel itself (Fig. 3-18b).

Thus, a researcher can monitor the progress of a protein purification procedure as the number of protein bands visible on the gel decreases after each new fractionation step. When compared with the positions to which proteins of known molecular weight migrate in the gel, the position of an unidentified protein can provide a good approximation of its molecular weight (Fig. 3-19). If the protein has two or more different subunits, generally the subunits are separated by the SDS treatment, and a separate band appears for each.

In this method, large proteins emerge from the column sooner than small ones — a somewhat counterintuitive result. The solid phase consists of cross-linked polymer beads with engineered pores or cavities of a particular size. Large proteins cannot enter the cavities and so take a shorter (and more rapid) path through the column, around the beads. Small proteins enter the cavities and are slowed by their more labyrinthine path through the column. Size-exclusion chromatography can also be used to approximate the size of a protein being purified, using methods similar to those described in Figure 3-19.

FIGURE 3-19 Estimating the molecular weight of a protein. The electrophoretic mobility of a protein on an SDS polyacrylamide gel is related to its molecular weight, Mr. (a) Standard proteins of known molecular weight are subjected to electrophoresis (lane 1). These marker proteins can be used to estimate the molecular weight of an unknown protein (lane 2). (b) A plot of log Mr of the marker proteins versus relative migration during electrophoresis is linear, which allows the molecular weight of the unknown protein to be read from the graph. (In similar fashion, a set of standard proteins with reproducible retention times on a size-exclusion column can be used to create a standard curve of retention time versus log Mr. The retention time of an unknown substance on the column can be compared with this standard curve to obtain an approximate Mr.)

Chromatographic methods are typically enhanced by the use of HPLC, or high-performance liquid chromatography. HPLC makes use of high-pressure pumps that speed the movement of the protein molecules down the column; it also uses higherquality chromatographic materials that can withstand the crushing force of the pressurized flow. By reducing the transit time on the column, HPLC can limit diffusional spreading of protein bands and thus can greatly improve resolution.

Mass spectrometry can provide a highly accurate measure of the molecular mass of a protein, readily distinguishing between single proton differences.

In 1988, two different techniques were introduced to permit transfer of macromolecules to the gas phase while limiting decomposition; these new capabilities revolutionized protein sequencing. In one technique, proteins are placed in a light-absorbing matrix. With a short pulse of laser light, the proteins are ionized and then desorbed from the matrix into the vacuum system. This process, known as matrix-assisted laser desorption/ionization mass spectrometry, or MALDI MS, is used to measure the mass of macromolecules.

A solution of analytes is passed through a charged needle that is kept at a high electrical potential, dispersing the solution into a fine mist of charged microdroplets. The solvent surrounding the macromolecules rapidly evaporates, leaving multiply charged macromolecular ions in the gas phase. This technique is called electrospray ionization mass spectrometry, or ESI MS. Protons added during passage through the needle give additional charge to the macromolecule. The m/z of the molecule can then be analyzed in the vacuum chamber. One method for analyzing m/z is called time of flight or TOF, in which ion acceleration in an electric field depends on m/z. A newer, moreefficient method is the Orbitrap, in which ions are trapped in orbit between an outer barrel-shaped electrode and an inner spindle electrode.

The process for determining the molecular mass of a protein with ESI MS is illustrated in Figure 3-28. As a protein is injected into the gas phase, it acquires a variable number of protons, and thus positive charges, from the solvent. The variable addition of these charges creates a spectrum of species with different mass-tocharge ratios. Each successive peak corresponds to a species that differs from that of its neighboring peak by a charge of 1 and a mass of 1 (one proton). The mass of the protein can be determined from any two neighboring peaks.

No generalizations can be made about the molecular weights of biologically active peptides and proteins in relation to their functions. Naturally occurring peptides range in length from two to many thousands of amino acid residues. Even the smallest peptides can have biologically important effects.

We can estimate the number of amino acid residues in a simple protein containing no other chemical constituents by dividing its molecular weight by 110. Although the average molecular weight of the 20 common amino acids is about 138, the smaller amino acids predominate in most proteins. If we take into account the proportions in which the various amino acids occur in an average protein (Table 3-1; the averages are determined by surveying the amino acid compositions of more than 1,000 different proteins), the average molecular weight of protein amino acids is nearer to 128. Because a molecule of water (Mr 18) is removed to create each peptide bond, the average molecular weight of an amino acid residue in a protein is about 128 − 18 = 110.

Hemoglobin, for example, has four polypeptide subunits: two identical α chains and two identical β chains, all four held together by noncovalent interactions. Each α subunit is paired in an identical way with a β subunit within the structure of this multisubunit protein, so that hemoglobin can be considered either a tetramer of four polypeptide subunits or a dimer of αβ protomers.

Tyrosine and tryptophan are significantly more polar than phenylalanine because of the tyrosine hydroxyl group and the nitrogen of the tryptophan indole ring. Tryptophan and tyrosine, and to a much lesser extent phenylalanine, absorb ultraviolet light (Fig. 36; see also Box 3-1). This accounts for the characteristic strong absorbance of light by most proteins at a wavelength of 280 nm, a property exploited by researchers in the characterization of proteins.

A wide range of biomolecules absorb light at characteristic wavelengths, just as tryptophan absorbs light at 280 nm (see Fig. 3-6). Measurement of light absorption by a spectrophotometer is used to detect and identify molecules and to measure their concentration in solution. The fraction of the incident light absorbed by a solution at a given wavelength is related to the thickness of the absorbing layer (path length) and the concentration of the absorbing species (Fig. 1). These two relationships are combined into the Lambert-Beer law,

However, with an absorbing layer of fixed path length, the absorbance, A, is directly proportional to the concentration of the absorbing solute. The molar extinction coefficient varies with the nature of the absorbing compound, the solvent, and the wavelength, and also with pH if the light-absorbing species is in equilibrium with an ionization state that has different absorbance properties.

To purify a protein, it is essential to have a way of detecting and quantifying that protein in the presence of many other proteins at each stage of the procedure. A common target of purification is one or another of the class of proteins called enzymes (Chapter 6). Each enzyme catalyzes a particular reaction that converts one biomolecule (the substrate) to another (the product). The amount of the protein in a given solution or tissue extract can be measured, or assayed, in terms of the catalytic effect the enzyme produces — that is, the increase in the rate at which its substrate is converted to reaction products when the enzyme is present.

The term activity refers to the total units of enzyme in a solution. The specific activity is the number of enzyme units per milligram of total protein (Fig. 3-22). The specific activity is a measure of enzyme purity: it increases during purification of an enzyme and becomes maximal and constant when the enzyme is pure (Table 3-5).

For proteins that are not enzymes, other quantification methods are required. Transport proteins can be assayed by their binding to the molecule they transport, and hormones and toxins by the biological effect they produce; for example, growth hormones will stimulate the growth of certain cultured cells. Some structural proteins represent such a large fraction of a tissue mass that they can be readily extracted and purified without a functional assay. The approaches are as varied as the proteins themselves.

The extraordinary binding affinity and specificity of antibodies make them valuable analytical reagents. Two types of antibody preparations are in use: polyclonal and monoclonal. Polyclonal antibodies are those produced by many different B lymphocytes responding to one antigen, such as a protein injected into an animal. Cells in the population of B lymphocytes produce antibodies that bind specific, different epitopes within the antigen. Thus, polyclonal preparations contain a mixture of antibodies that recognize different parts of the protein. Monoclonal antibodies, in contrast, are synthesized by a population of identical B cells (a clone) grown in cell culture. These antibodies are homogeneous, all recognizing the same epitope.

The specificity of antibodies has practical uses. In a versatile analytical technique, an antibody is attached to a reagent that makes it easy to detect (Fig. 5-25). When the antibody binds the target protein, the label reveals the presence of the protein in a solution or its location in a gel, or even in a living cell (Fig. 5-25a). In one application of this technique, an immunoblot or Western blot assay (Fig. 5-26b), proteins that have been separated by gel electrophoresis are transferred electrophoretically to a nitrocellulose membrane. After washing, the membrane is treated successively with primary antibody, secondary antibody linked to enzyme, and substrate. A colored precipitate forms only along the band containing the protein of interest. Immunoblotting allows the detection of a minor component in a sample and provides an approximation of its molecular weight.

FIGURE 5-25 Antibodies as analytical reagents. (a) A schematic representation of the use of the specific reaction of an antibody with its antigen to identify and quantify a specific protein in a complex sample. (b) One common application is the immunoblot. Lanes 1 to 3 are from an SDS gel; samples from successive stages in the purification of a protein kinase were separated and stained with Coomassie blue. Lanes 4 to 6 show the same samples, but these were electrophoretically transferred to a nitrocellulose membrane after separation on an SDS gel. The membrane was then “probed” with antibody against the protein kinase. The numbers between the SDS gel and the immunoblot indicate Mr in thousands.

The data are assembled in a purification table similar to Table 3-5. A protein is generally considered pure when further purification steps fail to increase specific activity and when only a single protein species can be detected (for example, by electrophoresis in the presence of SDS).

The fourth lane shows the proteins in a crude cellular extract. Subsequent lanes (left to right) show the proteins that are present after successive purification steps. Although the protein looks pure in lane 6, two more steps are needed to remove minor contaminants not evident on the gel. The purified protein is a single polypeptide chain (Mr~ 38,000), as seen in the rightmost lane.

FIGURE 3-20 Isoelectric focusing. This technique separates proteins according to their isoelectric points. A protein mixture is placed on a gel strip containing an immobilized pH gradient. With an applied electric field, proteins enter the gel and migrate until each reaches a pH that is equivalent to its pI. Remember that when pH = pI, the net charge of a protein is zero.

Chromatographic methods are typically enhanced by the use of HPLC, or high-performance liquid chromatography. HPLC makes use of high-pressure pumps that speed the movement of the protein molecules down the column; it also uses higherquality chromatographic materials that can withstand the crushing force of the pressurized flow. By reducing the transit time on the column, HPLC can limit diffusional spreading of protein bands and thus can greatly improve resolution.

In reality, however, each protein has a specific chemical or structural function, which suggests that each has a unique three-dimensional structure (Fig. 4-1). How stable is this structure, what factors guide its formation, and what holds it together? By the late 1920s, several proteins had been crystallized, including hemoglobin (Mr 64,500) and the enzyme urease (Mr 483,000).

However, the orderly bundles formed by fibrous proteins are not crystals — the molecules are aligned side by side, but not all are oriented in the same direction. More-detailed three-dimensional structural information about proteins requires a highly ordered protein crystal. The structures of many proteins are not yet known, simply because they have proved difficult to crystallize. Practitioners have compared making protein crystals to holding together a stack of bowling balls with cellophane tape.