第二章 氨基酸、肽和蛋白质

Proteins are polymers of amino acids, with each amino acid residue joined to its neighbor by a specific type of covalent bond. (The term “residue” reflects the loss of the elements of water when one amino acid is joined to another.) Proteins can be broken down (hydrolyzed) to their constituent amino acids by a variety of methods, and the earliest studies of proteins naturally focused on the free amino acids derived from them.

Twenty different amino acids are commonly found in proteins. The first to be discovered was asparagine, in 1806. The last of the 20 to be found, threonine, was not identified until 1938. All the amino acids have trivial or common names, in some cases derived from the source from which they were first isolated. Asparagine was first found in asparagus, and glutamate in wheat gluten; tyrosine was first isolated from cheese (its name is derived from the Greek tyros, “cheese”); glycine (Greek glykos, “sweet”) was so named because of its sweet taste.

Learning the names, structures, and chemical properties of the 20 common amino acids found in proteins is one of the key memorization trials of every beginning biochemistry student. The necessity rapidly becomes apparent in succeeding chapters. It is impossible to discuss protein structure, protein function, ligand-binding sites, enzyme active sites, and most other biochemical topics without this foundation. The amino acids are part of the biochemistry vocabulary.

All 20 of the common amino acids are α-amino acids. They have a carboxyl group and an amino group bonded to the same carbon atom (the α carbon) (Fig. 3-2). They differ from each other in their side chains, or R groups, which vary in structure, size, and electric charge, and which influence the solubility of the amino acids in water. In addition to these 20 amino acids, there are many less common ones.

The three-letter code is easily understood, the abbreviations generally consisting of the first three letters of the amino acid name. The one-letter code was devised by Margaret Oakley Dayhoff, considered by many to be the founder of the field of bioinformatics. The one-letter code reflects an attempt to reduce the size of the data files (in an era of limited computer memory) used to describe amino acid sequences. It was designed to be easily memorized, and understanding its origin can help students do just that.

For six amino acids (CHIMSV), the first letter of the amino acid name is unique and thus is used as the symbol. For five others (AGLPT), the first letter of the name is not unique but is assigned to the amino acid that is most common in proteins (for example, leucine is more common than lysine).

For all the common amino acids except glycine, the α carbon is bonded to four different groups: a carboxyl group, an amino group, an R group, and a hydrogen atom (Fig. 3-2; in glycine, the R group is another hydrogen atom). The α-carbon atom is thus a chiral center (p. 61). Because of the tetrahedral arrangement of the bonding orbitals around the α-carbon atom, the four different groups can occupy two unique spatial arrangements, and thus amino acids have two possible stereoisomers. Since they are nonsuperposable mirror images of each other (Fig. 3-3), the two forms represent a class of stereoisomers called enantiomers (see Fig. 1-21).

For most other organic molecules, carbon atoms are simply numbered from one end, giving highest priority (C-1) to the carbon with the substituent containing the atom of highest atomic number. Within this latter convention, the carboxyl carbon of an amino acid would be C-1 and the α carbon would be C-2.

For all chiral compounds, stereoisomers having a configuration related to that of L-glyceraldehyde are designated L, and stereoisomers related to D-glyceraldehyde are designated D. The functional groups of L- alanine are matched with those of L-glyceraldehyde by aligning those that can be interconverted by simple, one-step chemical reactions. Thus the carboxyl group of L-alanine occupies the same position about the chiral carbon as does the aldehyde group of L-glyceraldehyde, because an aldehyde is readily converted to a carboxyl group via a one-step oxidation.

However, not all L-amino acids are levorotatory, and the convention shown in Figure 3-4 was needed to avoid potential ambiguities about absolute configuration. By Fischer’s convention, L and D refer only to the absolute configuration of the four substituents around the chiral carbon, not to optical properties of the molecule.

Nearly all biological compounds with a chiral center occur naturally in only one stereoisomeric form, either D or L. The amino acid residues in protein molecules are almost all L stereoisomers, with less than 1% being found in the D-configuration.

It is remarkable that virtually all amino acid residues in proteins are L stereoisomers. When chiral compounds are formed by ordinary chemical reactions, the result is a racemic mixture of D and L isomers, which are difficult for a chemist to distinguish and separate.

Only the L stereoisomers of amino acids, with a configuration related to the absolute configuration of the reference molecule L-glyceraldehyde, are found in proteins. Amino acids can be classified into five types on the basis of the polarity and charge (at pH 7) of their R groups.

Knowledge of the chemical properties of the common amino acids is central to an understanding of biochemistry. The topic can be simplified by grouping the amino acids into five main classes based on the properties of their R groups (Table 3-1), particularly their polarity, or tendency to interact with water at biological pH (near pH 7.0). The polarity of the R groups varies widely, from nonpolar and hydrophobic (water-insoluble) to highly polar and hydrophilic (water-soluble). A few amino acids — especially glycine, histidine, and cysteine — are somewhat difficult to characterize or do not fit perfectly in any one group. They are assigned to particular groupings based on considered judgments rather than absolutes.

Nonpolar, Aliphatic R Groups The R groups in this class of amino acids are nonpolar and hydrophobic. The side chains of alanine, valine, leucine, and isoleucine tend to cluster together within proteins, stabilizing protein structure through the hydrophobic effect. Glycine has the simplest structure. Although it is most easily grouped with the nonpolar amino acids, its very small side chain makes no real contribution to interactions driven by the hydrophobic effect. Methionine, one of the two sulfur-containing amino acids, has a slightly nonpolar thioether group in its side chain. Proline has an aliphatic side chain with a distinctive cyclic structure. The secondary amino (imino) group of proline residues is held in a rigid conformation that reduces the structural flexibility of polypeptide regions containing proline.

Aromatic R Groups Phenylalanine, tyrosine, and tryptophan, with their aromatic side chains, are relatively nonpolar (hydrophobic). All can contribute to the hydrophobic effect. The hydroxyl group of tyrosine can form hydrogen bonds, and it is an important functional group in some enzymes.

Polar, Uncharged R Groups The R groups of these amino acids are more soluble in water, or more hydrophilic, than those of the nonpolar amino acids because they contain functional groups that form hydrogen bonds with water. This class of amino acids includes serine, threonine, cysteine, asparagine, and glutamine. The polarity of serine and threonine is contributed by their hydroxyl groups, and the polarity of asparagine and glutamine is contributed by their amide groups. Cysteine is an outlier here because its polarity, contributed by its sulfhydryl group, is quite modest. Cysteine is a weak acid and can make weak hydrogen bonds with oxygen or nitrogen.

Asparagine and glutamine are the amides of two other amino acids also found in proteins —aspartate and glutamate, respectively — to which asparagine and glutamine are easily hydrolyzed by acid or base. Cysteine is readily oxidized to form a covalently linked dimeric amino acid called cystine, in which two cysteine molecules or residues are joined by a disulfide bond (Fig. 3-7). The disulfide-linked residues are strongly hydrophobic (nonpolar). Disulfide bonds play a special role in the structures of many proteins by forming covalent links between parts of a polypeptide molecule or between two different polypeptide chains.

Positively Charged (Basic) R Groups The most hydrophilic R groups are those that are either positively charged or negatively charged. The amino acids in which the R groups have significant positive charge at pH 7.0 are lysine, which has a second primary amino group at the ε position on its aliphatic chain; arginine, which has a positively charged guanidinium group; and histidine, which has an aromatic imidazole group. As the only common amino acid having an ionizable side chain with pKa near neutrality, histidine may be positively charged (protonated form) or uncharged at pH 7.0. His residues facilitate many enzyme-catalyzed reactions by serving as proton donors/acceptors. Negatively Charged (Acidic) R Groups The two amino acids having R groups with a net negative charge at pH 7.0 are aspartate and glutamate, each of which has a second carboxyl group.

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,

where I0 is the intensity of the incident light, I is the intensity of the transmitted light, the ratio I/I0 (the inverse of the ratio in the equation) is the transmittance, ε is the molar extinction coefficient (in units of liters per mole-centimeter), c is the concentration of the absorbing species (in moles per liter), and l is the path length of the light-absorbing sample (in centimeters). The Lambert-Beer law assumes that the incident light is parallel and monochromatic (of a single wavelength) and that the solvent and solute molecules are randomly oriented. The expression log (I0/I) is called the absorbance, designated A.

It is important to note that each successive millimeter of path length of absorbing solution in a 1.0 cm cell absorbs not a constant amount but a constant fraction of the light that is incident upon it. 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.

In addition to the 20 common amino acids, proteins may contain residues created by modification of common residues already incorporated into a polypeptide — that is, through postsynthetic modification (Fig. 3-8a). Among these uncommon amino acids are 4-hydroxyproline, a derivative of proline found in the fibrous protein collagen

Selenocysteine contains selenium rather than the sulfur of cysteine. Actually derived from serine, selenocysteine is a constituent of just a few known proteins. Pyrrolysine is found in a few proteins in several methanogenic (methane-producing) archaea and in one known bacterium; it plays a role in methane biosynthesis.

Some amino acid residues in a protein may be modified transiently to alter the protein’s function. The addition of phosphoryl, methyl, acetyl, adenylyl, ADP-ribosyl, or other groups to particular amino acid residues can increase or decrease a protein’s activity (Fig. 3-8b). Phosphorylation is a particularly common regulatory modification. Covalent modification as a protein regulatory strategy is discussed in more detail in Chapter 6.

Some 300 additional amino acids have been found in cells. They have a variety of functions, but not all are constituents of proteins. Ornithine and citrulline (Fig. 3-8c) deserve special note because they are key intermediates (metabolites) in the biosynthesis of arginine (Chapter 22) and in the urea cycle (Chapter 18).

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).

Acid-base titration involves the gradual addition or removal of protons (Chapter 2). Figure 3-10 shows the titration curve of the diprotic form of glycine. The two ionizable groups of glycine, the carboxyl group and the amino group, are titrated with a strong base such as NaOH. The plot has two distinct stages, corresponding to deprotonation of two different groups on glycine.

In the first stage of the titration, the —COOH group of glycine (with its lower pKa) loses its proton. At the midpoint of this stage, equimolar concentrations of the proton-donor (+H3N—CH2—COOH) and the proton-acceptor (+H3N—CH2—COO−) species are present. As in the titration of any weak acid, a point of inflection is reached at this midpoint where the pH is equal to the pKa of the protonated group that is being titrated (see Fig. 2-17). For glycine, the pH at the midpoint is 2.34; thus its —COOH group has a pKa (labeled pK1 in Fig. 3-10) of 2.34.

As the titration of glycine proceeds, another point of inflection is reached at pH 5.97; at this point, removal of the first proton is essentially complete and removal of the second has just begun. At this pH, glycine is present largely as the dipolar ion (zwitterion) +H3N—CH2—COO−. We shall return to the significance of this inflection point in the titration curve (labeled pI in Fig. 3-10) shortly.

Note that the carboxyl group of glycine is over 100 times more acidic (more easily ionized) than the carboxyl group of acetic acid, which, as we saw in Chapter 2, has a pKa of 4.76 — about average for a carboxyl group attached to an otherwise unsubstituted aliphatic hydrocarbon. The perturbed pKa of glycine is caused primarily by the nearby positively charged amino group on the α-carbon atom, an electronegative group that tends to pull electrons toward it (a process called electron withdrawal), as described in Figure 3-11. The opposite charges on the resulting zwitterion are also somewhat stabilizing.

Similarly, the pKa of the amino group in glycine is perturbed downward relative to the average pKa of an amino group. This effect is due largely to electron withdrawal by the electronegative oxygen atoms in the carboxyl groups, increasing the tendency of the amino group to give up a proton. Hence, the α-amino group has a pKa that is lower than that of an aliphatic amine such as methylamine (Fig. 3-11). In short, the pKa of any functional group is greatly affected by its chemical environment, a phenomenon sometimes exploited in the active sites of enzymes to promote exquisitely adapted reaction mechanisms that depend on the perturbed pKa values of proton donor/acceptor groups of specific residues.

The second piece of information provided by the titration curve of glycine is that this amino acid has two regions of buffering power. One of these is the relatively flat portion of the curve, extending for approximately 1 pH unit on either side of the first pKa of 2.34, indicating that glycine is a good buffer near this pH. The other buffering zone is centered around pH 9.60. (Note that glycine is not a good buffer at the pH of intracellular fluid or blood, about 7.4.) Within the buffering ranges of glycine, the Henderson-Hasselbalch equation (p. 60) can be used to calculate the proportions of proton-donor and proton-acceptor species of glycine required to make a buffer at a given pH.

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. For glycine, which has no ionizable group in its side chain, the isoelectric point is simply the arithmetic mean of the two pKa values:

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. At pH 1.0, for example, glycine exists almost entirely as the form +H3N—CH2—COOH with a net positive charge of 1.0. At pH 2.34, where there is an equal mixture of +H3N—CH2—COOH and +H3N—CH2—COO−, the average or net positive charge is 0.5. The sign and the magnitude of the net charge of any amino acid at any pH can be predicted in the same way.

The shared properties of many amino acids permit some simplifying generalizations about their acid-base behaviors. First, all amino acids with a single α-amino group, a single α-carboxyl group, and an R group that does not ionize have titration curves resembling that of glycine (Fig. 3-10).

Second, amino acids with an ionizable R group have more complex titration curves, with three stages corresponding to the three possible ionization steps; thus, they have three pKa values. The additional stage for the titration of the ionizable R group merges to some extent with that for the titration of the α-carboxyl group, the titration of the α-amino group, or both. The titration curves for two amino acids of this type, glutamate and histidine, are shown in Figure 3-12.

The isoelectric points reflect the nature of the ionizing R groups that are present. For example, glutamate has a pI of 3.22, considerably lower than that of glycine. This is due to the presence of two carboxyl groups, which, at the average of their pKa values (3.22), contribute a net charge of −1 that balances the +1 contributed by the amino group. Similarly, the pI of histidine, with two groups that are positively charged when protonated, is 7.59 (the average of the pKa values of the amino and imidazole groups), much higher than that of glycine.

Amino acids vary in their acid-base properties and have characteristic titration curves. Monoamino monocarboxylic amino acids (with nonionizable R groups) are diprotic acids (+H3NCH(R)COOH) at low pH and exist in several different ionic forms as the pH is increased. Amino acids with ionizable R groups have additional ionic species, depending on the pH of the medium and the pKa of the R group.

Two amino acid molecules can be covalently joined through a substituted amide linkage, termed a peptide bond, to yield a dipeptide. Such a linkage is formed by removal of the elements of water (dehydration) — a hydroxyl moiety from the αcarboxyl group of one amino acid and a hydrogen atom the αamino group of another (Fig. 3-13).

Peptide bond formation is an example of a condensation reaction, a common class of reactions in living cells. The reverse reaction, bond breakage involving water, is an example of hydrolytic cleavage or hydrolysis. Under standard biochemical conditions, the equilibrium for the reaction shown in Figure 3-13 favors the hydrolysis of the dipeptide into amino acids. To make the condensation reaction thermodynamically more favorable, the carboxyl group must be chemically modified or activated so that the hydroxyl group can be more readily eliminated. A chemical approach to this problem is outlined later in this chapter. The biological approach to peptide bond formation is a major topic of Chapter 27.

Three amino acids can be joined by two peptide bonds to form a tripeptide; similarly, four amino acids can be linked to form a tetrapeptide, five to form a pentapeptide, and so forth. When a few amino acids are joined in this fashion, the structure is called an oligopeptide. When many amino acids are joined, the product is called a polypeptide. Proteins may have thousands of amino acid residues.

Figure 3-14 shows the structure of a pentapeptide. In a peptide, the amino acid residue at the end with a free α-amino group is the amino-terminal (or N-terminal) residue; the residue at the other end, which has a free carboxyl group, is the carboxylterminal (C-terminal) residue.

Although hydrolysis of a peptide bond is an exergonic reaction, it occurs only slowly because it has a high activation energy (p. 25). As a result, the peptide bonds in proteins are quite stable, with an average half-life (t1/2) of about 7 years under most intracellular conditions.

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).

When an amino acid becomes a residue in a peptide, its chemical environment is altered, and the pKa value for an ionizable R group can change somewhat. The pKa values for R groups listed in Table 3-1 can be a useful guide to the pH range in which a given group will ionize, but they cannot be strictly applied when an amino acid becomes part of a peptide.

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. Consider the commercially synthesized dipeptide L-aspartyl-L-phenylalanine methyl ester, the artificial sweetener better known as aspartame or NutraSweet.

Many small peptides exert their effects at very low concentrations. For example, a number of vertebrate hormones (Chapter 23) are small peptides. These include oxytocin (nine amino acid residues), which is secreted by the posterior pituitary gland and stimulates uterine contractions, and thyrotropinreleasing factor (three residues), which is formed in the hypothalamus and stimulates the release of another hormone, thyrotropin, from the anterior pituitary gland. Some extremely toxic mushroom poisons, such as amanitin, are also small peptides, as are many antibiotics.

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. A few proteins contain two or more polypeptide chains linked covalently. For example, the two polypeptide chains of insulin are linked by disulfide bonds. In such cases, the individual polypeptides are not considered subunits; instead they are commonly referred to simply as chains.

The amino acid composition of proteins is also highly variable. The 20 common amino acids almost never occur in equal amounts in a protein. Some amino acids may occur only once or not at all in a given type of protein; others may occur in large numbers. Table 3-3 shows the amino acid composition of bovine cytochrome c and chymotrypsinogen, the inactive precursor of the digestive enzyme chymotrypsin. These two proteins, with very different functions, also differ significantly in the relative numbers of each kind of amino acid residue.

In some common analyses, such as acid hydrolysis, Asp and Asn are not readily distinguished from each other and are together designated Asx (or B). Similarly, when Glu and Gln cannot be distinguished, they are together designated Glx (or Z). In addition, Trp is destroyed by acid hydrolysis. Additional procedures must be employed to obtain an accurate assessment of complete amino acid content.

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.

Many proteins — for example, the enzymes ribonuclease A and chymotrypsin — contain only amino acid residues and no other chemical constituents. However, some proteins contain permanently associated chemical components in addition to amino acids; these are called conjugated proteins.

Conjugated proteins are classified on the basis of the chemical nature of their prosthetic groups (Table 3-4); for example, lipoproteins contain lipids, glycoproteins contain sugar groups, and metalloproteins contain a specific metal. Some proteins contain more than one prosthetic group. Usually the prosthetic group plays an important role in the protein’s biological function.

Amino acids can be joined covalently through peptide bonds to form peptides and proteins. Cells generally contain thousands of different proteins, each with a different biological activity. The ionization behavior of peptides reflects their ionizable side chains as well as the terminal α-amino and α-carboxyl groups. Proteins can be very long polypeptide chains of 100 to several thousand amino acid residues. However, some naturally occurring peptides have only a few amino acid residues. Some proteins are composed of several noncovalently associated polypeptide chains, called subunits. Simple proteins yield only amino acids on hydrolysis; conjugated proteins contain in addition some other component, such as a metal or organic prosthetic group.

A description of all covalent bonds (mainly peptide bonds and disulfide bonds) linking amino acid residues in a polypeptide chain is its primary structure. The most important element of primary structure is the sequence of amino acid residues. Secondary structure refers to particularly stable arrangements of amino acid residues giving rise to recurring structural patterns. Tertiary structure describes all aspects of the three-dimensional folding of a polypeptide. When a protein has two or more polypeptide subunits, their arrangement in space is referred to as quaternary structure. Our exploration of proteins will eventually include complex protein machines consisting of dozens to thousands of subunits. Primary structure is the focus of the remainder of this chapter; we discuss the higher levels of structure in Chapter 4.

In both species, each type of protein has a unique amino acid sequence that confers a particular threedimensional structure. This structure in turn confers a unique function. Amino acid sequences are important elements of the broader realm of biological information. They are a major functional expression of information stored in DNA in the form of genes. The sequences are not at all random. Each protein has a distinctive number and sequence of amino acid residues.

Some simple observations illustrate the functional importance of primary structure, or the amino acid sequence of a protein. First, as we have already noted, proteins with different functions always have different amino acid sequences. Second, thousands of human genetic diseases have been traced to the production of proteins with less activity or altered activity. The alteration can range from a single change in the amino acid sequence (as in sickle cell disease, described in Chapter 5) to deletion of a larger portion of the polypeptide chain (as in most cases of Duchenne muscular dystrophy: a large deletion in the gene encoding the protein dystrophin leads to production of a shortened, inactive protein).

The amino acid sequence for a particular protein is not absolutely fixed, or invariant. Virtually all of the proteins in humans are polymorphic, having amino acid sequence variants in the human population. Many human proteins are polymorphic even within an individual, with amino acid variations occurring due to processes that will be described in Part III of this text. Some of these variations have little or no effect on the function of the protein; others may affect function dramatically. Furthermore, proteins that carry out a broadly similar function in distantly related species can differ greatly in overall size and amino acid sequence.

Although the amino acid sequence in some regions of the primary structure might vary considerably without affecting biological function, most proteins contain crucial regions that are essential to their function and thus have sequences that are conserved. The fraction of the overall sequence that is critical varies from protein to protein, complicating the task of relating sequence to three-dimensional structure, and structure to function. Before we can consider this problem further, however, we must examine how sequence information is obtained.

However, an array of techniques derived from traditional methods of polypeptide sequencing made important contributions to the broader field of protein chemistry. The method used by Sanger to sequence insulin is based on the classical method for direct chemical sequencing of proteins from the amino terminus, the two-step Edman degradation developed by Pehr Edman.

The sequence of a protein can be predicted from the sequence of the gene encoding it, which is usually available in genomic databases. Direct sequencing can also be provided by mass spectrometry. Many methods used in traditional protein sequencing protocols remain valuable for labeling proteins or breaking them into parts for functional and structural analysis.

For example, the amino-terminal α-amino group of a protein can be labeled with 1-fluoro-2,4-dinitrobenzene (FDNB), dansyl chloride, or dabsyl chloride (Fig. 3-25). These reagents also label the ε-amino group of lysine residues. Disulfide bonds within a polypeptide or between polypeptide subunits can be broken irreversibly (Fig. 3-26).

Enzymes called proteases catalyze the hydrolytic cleavage of peptide bonds and provide the most common method to break a protein into parts. Some proteases cleave only the peptide bond adjacent to particular amino acid residues (Table 3-6) and thus fragment a polypeptide chain in a predictable and reproducible way. A few chemical reagents also cleave the peptide bond adjacent to specific residues. Among proteases, the digestive enzyme trypsin catalyzes the hydrolysis of only those peptide bonds in which the carbonyl group is contributed by either a Lys or an Arg residue, regardless of the length or amino acid sequence of the chain. A polypeptide with three Lys and/or Arg residues will usually yield four smaller peptides upon cleavage with trypsin. Moreover, all except one of these will have a carboxyl-terminal Lys or Arg.

The capacity to modify proteins in specific ways has many applications in the lab. The methods used to break disulfide bonds can also be used to denature proteins when that is required. The development of reagents to label the aminoterminal amino acid residue led eventually to the development of an array of reagents that could react with specific groups at many locations on a protein. For example, the sulfhydryl group on Cys residues can be modified with iodoacetamides, maleimides, benzyl halides, and bromomethyl ketones (Fig. 3-27).

Unknown purified proteins can be identified, and their mass can be accurately determined. When coupled to powerful peptide separation protocols, mass spectrometry can document a complete cellular proteome —defined as the entire complement of proteins in a cell, including estimates of their relative abundance — in just an hour.

The mass spectrometer has been an indispensable tool in chemistry for more than a century. Molecules to be analyzed, referred to as analytes, are first ionized in a vacuum. When the newly charged molecules are introduced into an electric and/or magnetic field, their paths through the field are a function of their mass-to-charge ratio, m/z. This measured property of the ionized species can be used to deduce the mass (m) of the analyte with very high precision.

As the m/z measurements are made in the gas phase, the technique was long limited to relatively small molecules. 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 trajectory of the electrons, related to their mass and charge, is detected and converted to m/z by a Fourier transform.

The mixture is then injected into a mass spectrometer that has two mass filters in tandem (Fig. 329a, top). In the first, the peptide mixture is sorted so that only one of the several types of peptides produced by cleavage emerges at the other end. The sample of the selected peptide, each molecule of which has a charge somewhere along its length, then travels through a vacuum chamber between the two mass spectrometers. In this collision cell, the peptide is further fragmented by high-energy impact with a “collision gas” such as helium or argon that is bled into the vacuum chamber. Each individual peptide is broken in only one place, on average. Although the breaks are not hydrolytic, most occur at the peptide bonds.

The second mass filter then measures the m/z ratios of all the charged fragments. This process generates one or more sets of peaks. A given set of peaks (Fig. 3-29b) consists of all the charged fragments that were generated by breaking the same type of bond (but at different points in the peptide). One set of peaks includes only the fragments in which the charge was retained on the amino-terminal side of the broken bonds; another includes only the fragments in which the charge was retained on the carboxylterminal side of the broken bonds. Each successive peak in a given set has one less amino acid than the peak before. The difference in mass from peak to peak identifies the amino acid that was lost in each case, thus revealing the sequence of the peptide. The only ambiguities involve leucine and isoleucine, which have the same mass.

The organism of interest is generally one in which the genomic sequence is known. Cellular proteins are first isolated in an extract, then digested into relatively short peptides by a protease such as trypsin. The very complex mixture of peptides is subjected to chromatography, so that resolved peptides are introduced to the mass spectrometer successively. Transfer from the liquid phase to the gas phase is facilitated by MALDI or ESI. Each peptide is analyzed for amino acid sequence, and that sequence is compared to the known genomic sequence available in databases to identify the protein it came from. Because more peptides are generated from the more common proteins in the mixture, the exercise also provides a measure of protein abundance.

Many peptides are potentially useful as pharmacologic agents, and their production is of considerable commercial importance. In addition to its commercial applications, the synthesis of specific peptide portions of larger proteins is an increasingly important tool for the study of protein structure and function.

The major breakthrough in this technology was provided by R. Bruce Merrifield in 1962. His innovation was to synthesize a peptide while keeping one end attached to a solid support. The support is an insoluble polymer (resin) contained within a column, similar to that used for chromatographic procedures. The peptide is built up on this support one amino acid at a time, through a standard set of reactions in a repeating cycle (Fig. 330). At each successive step in the cycle, protective chemical groups block unwanted reactions.

The technology for chemical peptide synthesis has been automated. An important limitation of the process is the efficiency of each chemical cycle. Incomplete reaction at one stage can lead to formation of an impurity (in the form of a shorter peptide) in the next. The chemistry has been optimized to permit the synthesis of proteins of 100 amino acid residues in a few days in reasonable yield. A very similar approach is used to synthesize nucleic acids (see Fig. 8-33). It is worth noting that this technology, impressive as it is, still pales when compared with biological processes. The same 100-residue protein would be synthesized with exquisite fidelity in about 5 seconds in a bacterial cell.

Methods for the efficient ligation (joining together) of peptides allow the assembly of synthetic peptides into larger polypeptides and proteins. Novel forms of proteins can be created with precisely positioned chemical groups, including those that might not normally be found in a cellular protein. This provides one approach to test theories of enzyme catalysis, to create proteins with altered chemical properties, and to design protein sequences that will fold into particular structures. This last application provides the ultimate test of our ability to relate the primary structure of a peptide to the three-dimensional structure that it takes up in solution.

We do not understand in detail exactly how the amino acid sequence determines three-dimensional structure, nor can we always predict function from sequence. However, protein families that have some shared structural or functional features can be readily identified on the basis of amino acid sequence similarities. Individual proteins are assigned to families based on the degree of similarity in amino acid sequence. Members of a family are usually identical across 25% or more of their sequences, and proteins in these families generally share at least some structural and functional characteristics. Some families, however, are defined by identities involving only a few amino acid residues that are critical to a certain function.

The residues that are less important to function may vary over time — that is, one amino acid may substitute for another — and these variable residues can provide the information to trace evolution. Some proteins have more variable amino acid residues than others. For these and other reasons, different proteins evolve at different rates.

Another complicating factor in tracing evolutionary history is the rare transfer of a gene or a group of genes from one organism to another, a process called horizontal gene transfer. The transferred genes may be similar to the genes they were derived from in the original organism, whereas most other genes in the two organisms may be only distantly related. An example of horizontal gene transfer is the recent rapid spread of antibioticresistance genes in bacterial populations. The proteins derived from these transferred genes would not be good candidates for the study of bacterial evolution, because they share only a very limited evolutionary history with their “host” organisms.

For example, a protein called EF-1α (elongation factor 1α) is involved in the synthesis of proteins in all eukaryotes. A similar protein, EF-Tu, with the same function, is found in bacteria. Similarities in sequence and function indicate that EF-1α and EF-Tu are members of a family of proteins that share a common ancestor. The members of protein families are called homologous proteins, or homologs. The concept of a homolog can be further refined. If two proteins in a family (that is, two homologs) are present in the same species, they are referred to as paralogs. Homologs from different species are called orthologs. The process of tracing evolution involves first identifying suitable families of homologous proteins and then using them to reconstruct evolutionary paths.

A more useful analysis also considers the chemical properties of substituted amino acids. Many of the amino acid differences within a protein family may be conservative — that is, an amino acid residue is replaced by a residue that has similar chemical properties. For example, a Glu residue may substitute in one family member for the Asp residue found in another; both amino acids are negatively charged. Logically, such a conservative substitution should receive a higher score in a sequence alignment than a nonconservative substitution does — for example, in replacement of the Asp residue with a hydrophobic Phe residue.

To use a protein family to explore evolution, researchers identify family members with similar molecular functions in the widest possible range of organisms. The sequence divergence in these protein families allows segregation of organisms into classes based on their evolutionary relationships. Certain segments of a protein sequence may be found in the organisms of one taxonomic group but not in other groups; these segments can be used as signature sequences for the group in which they are found. An example of a signature sequence is an insertion of 12 amino acids near the amino terminus of the EF-1α/EF-Tu proteins in all archaea and eukaryotes but not in bacteria (Fig. 332). This particular signature is one of many biochemical clues that can help establish the evolutionary relatedness of eukaryotes and archaea.

Most amino acid sequences are deduced from genomic sequences and by mass spectrometry. Methods derived from classical approaches to protein sequencing remain important in protein chemistry. Short proteins and peptides (up to about 100 residues) can be chemically synthesized. The peptide is built up, one amino acid residue at a time, while tethered to a solid support. Protein sequences are a rich source of information about protein structure and function. Bioinformatics can analyze changes in the amino acid sequences of homologous proteins over time to trace the evolution of life on Earth.

Biochemists’ understanding of protein structure and function has been derived from the study of many individual proteins. To study a protein in detail, the researcher must be able to separate it from other proteins in pure form and must have the techniques to determine its properties. The necessary methods come from protein chemistry, a discipline as old as biochemistry itself and one that retains a central position in biochemical research.

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). Once the extract or organelle preparation is ready, various methods are available for purifying one or more of the proteins it contains.

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. Low-speed centrifugation is then used to remove the precipitated proteins from those remaining in solution.

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 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.

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.

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.

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?

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.

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

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.

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.

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.

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.

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.

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.

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.

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.

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).

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.

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).

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 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%).

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.