Peptide Bond

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Peptide Bonds Link Amino Acids Together in Proteins

Proteins are formed as a linear, unbranched chain of amino acids. The amino acids are covalently linked by a peptide bond formed between the amino group of one amino acid and the carboxyl group of the next. The formation of the peptide bond is a dehydration reaction, as shown in Figure 2.3.8.

Figure 2.3.8. Formation of a peptide bond between two amino acids. The overall reaction is shown. The actual reaction involves many intermediary steps, catalyzed by enzymes.

Cells make proteins by the sequential addition of amino acids to the carboxyl terminus of the growing chain. This is accomplished on the ribosome when the amino acids bound to specific tRNAs (transfer RNAs) bind to the appropriate triplet codon of the mRNA. The amino acid is attached to the tRNA via its carboxyl group. When the peptide bond is formed, the entire growing chain is transferred to the free amino terminal of the next amino acid. The ribosome then moves one frame (nucleotide triplet) and the process is repeated (see Chapter 2.2).

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Chapter

Abstract

A peptide or amide (–CO–NH–) bond is formed by the linking of the carboxyl group of one amino acid with the amino group of another with the loss of a water molecule. In this bond, the oxygen atom of the carbonyl group is in the trans position with respect to the hydrogen on the nitrogen atom and all of these four atoms are planar due to resonance. This bond has a partially (40%) double-bond character, and the half-life for the hydrolysis of peptide bonds is 350–600 years at room temperature and at pH 4–8. The selective hydrolysis of the peptide bond of peptides and proteins is required in a wide range of biological, biotechnological, and industrial applications.

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5.2.2 Peptide bond

Chemically, a covalent bond can be formed between the carboxyl group of one amino acid and the amino group of another by the loss of a water molecule. This is called an amide bond or a peptide bond (Fig. 5.13). The bond formation can be reversed by the addition of water (hydrolysis). The equilibrium of the reaction is more toward hydrolysis than synthesis. Hence, an input of free energy is required for its biosynthesis. In the cell, peptide bond formation is an enzymatically controlled process (discussed in Chapter 11).

Fig. 5.13. Peptide bond formation. Upper panel: chemical structure; lower panel: three-dimensional structure. Coloring scheme for atoms as in Fig. 5.6.

The nature of the peptide bond is influenced by the phenomenon of resonance where electrons are delocalized over multiple atoms and a given molecule cannot be represented by a single valence structure. Two resonance structures are possible for the peptide bond (Fig. 5.14A). Due to resonance, the peptide bond possesses ~ 40% double-bond character which prevents rotation about this bond. As a consequence, the peptide group has a rigid, planar structure (Fig. 5.14B).

Fig. 5.14. The peptide bond. (A) Resonance structure; (B) peptide plane.

A peptide group usually adopts the trans conformation with successive Cα atoms being positioned on opposite sides of the peptide bond joining them. This trans peptide group forms the basic repeating unit of the polypeptide backbone. We can see that, unlike the peptide bond, the other two bonds in the unit, the N-Cα and Cα-C bonds, are pure single bonds. Two adjacent planar peptide groups are free to rotate about these bonds, provided there is no steric hindrance (Fig. 5.15). The two rotations are described by torsion angles (also called dihedral angles)—the rotation around N-Cα being designated as phi (ϕ), while that around Cα-C as psi (ψ). The backbone of a polypeptide can be considered as a linked sequence of planar peptide groups, with rotatable covalent bonds alternating with rigid ones (Fig. 5.16).

Fig. 5.15. Rotations of adjacent planar peptide groups.

Fig. 5.16. Extended conformation of a polypeptide with the backbone shown as a linked sequence of planar peptide groups.

As already indicated, several (ϕ, ψ) combinations are restricted by the principle of steric exclusion. According to this principle, two atoms cannot be present in the same place at the same time. They cannot be closer to each other than the corresponding van der Waals distance. As a consequence, only certain values of ϕ and ψ are permitted. This is represented by a two-dimensional plot known as a Ramachandran diagram (named after its inventor, G.N. Ramachandran) (Fig. 5.17).

Fig. 5.17. A Ramachandran diagram. The shaded regions indicate the sterically permitted values of ϕ and ψ—the most favorable and borderline regions are shown in darker and lighter shades, respectively.

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Peptides

Amino acids can establish covalent bonds between the carboxyl group of one amino acid and the α amine group of another. This amide-type link is called a peptide bond and it is accompanied by the loss of water (Fig. 3.7).

Fig. 3.7. Peptide bond.

The product formed by linking two amino acids together is called a dipeptide. The subsequent binding of additional amino acid units to this dipeptide via peptide bonding generates tripeptides, tetrapeptides, pentapeptides, etc. The polymers formed by more than 10 amino acids are designated polypeptides. A polypeptide chain is considered a protein when it has a molecular mass greater than 6000 Daltons (Da). A Da is the unit of atomic mass; it is 1/12 the mass of one atom of 12C. Frequently the expression kilodalton (kDa), 1000 Da is used. Relative mass is the ratio between the molecular mass of a given substance and the mass of one atom of 12C, which corresponds to the mass of a molecule greater than 50 amino acids. Below this mass, the compounds are designated peptides. There is no precise distinction between peptides and proteins; 6000 Da is arbitrary and was chosen because it is the approximate mass of insulin, a hormone produced in the pancreas, which was the first protein for which the entire structure was deciphered.

At one end of every polypeptide chain, there is an amino acid with a free α-amine group. By convention, this end is considered the beginning of the chain and is called the amino-terminal or N-terminal portion of the polypeptide. The other end of the chain, containing the free carboxyl group, is considered the C-terminal end of the polypeptide chain.

When integrated into the peptide or protein chain, amino acids lose the H of the amine group and the OH of the carboxyl group that is involved in the peptide binding. The amino acid units forming the polymer are referred to as amino acid residues.

Nomenclature

Peptides are named following the order of the amino acids that constitute them, beginning from the N-terminal of the chain. Amino acid residues are indicated by the root of its name followed by the suffix “yl.” The last residue (with the free carboxyl group) is mentioned by its full name. For example, in the case of the hexapeptide composed of serine, aspartic acid, tyrosine, lysine, alanine, and cysteine (Fig. 3.8), the name is: seryl-tyrosyl-aspartyl-lysyl-alanyl-cysteine, which can be abbreviated: ser-asp-tyr-lys-ala-cys or SDYKAC.

Fig. 3.8. Hexapeptide seryl-tyrosyl-aspartyl-lysyl-alanyl-cysteine. The peptide bonds (red) form the backbone of the molecule.

Acid-base properties

The carboxyl and amine groups involved in the peptide bonds lose OH and H respectively and cannot ionize. Therefore, the acid-base properties of peptides are determined by the terminal amine and carboxyl groups as well as the ionizable groups in the side chains of the amino acid residues (COO−, ▬NH3−, SH). The pH of the medium influences the magnitude and sign of the net charge of a peptide in a similar manner to that described for amino acids. Peptides also have an isoelectric point, the pH in which there is an equal number of positive and negative charges in the molecule.

Biologically important peptides

Peptides have important functions, both in plants and animals. Generally, peptides are chains of amino acids linked by peptide bonds; however, some of them have peculiar characteristics, such as atypical peptide bonds, unusual amino acids or amino acid derivatives, and the formation of cyclic structures.

The number and type of amino acids, their sequence, spatial arrangement, and their biological activity are important determinants of the functional role of peptides. Glutathione is a peptide widely distributed in nature, found in bacteria, plants, and animals. This tripeptide is composed of glutamic acid, cysteine, and glycine. Glutamic acid is bound to the amine group of cysteine by an atypical peptide bond, which involves the distal or α-carboxyl group of glutamic acid.

When oxidized, glutathione forms a disulfide bridge (SS) with another molecule of glutathione in a reaction that is reversible. Glutathione participates in enzymatic redox systems. It contributes to the prevention of oxidative damage in blood cells and tissues.

Many hormones are peptides. Table 3.2 presents a list of some of them. Another interesting group of peptides are the enkephalins, which are pentapeptides released by the central nervous system, which are involved in nociception. They produce analgesia by binding to specific brain receptors of the family of the opioid receptors. Some peptides and compounds that contain a peptide as part of their molecule work as antibiotics, which are substances synthesized by microorganisms that have toxic effects on other organisms. They have an antimicrobial effect by disrupting the membrane of bacteria. Some of these antibiotics have a cyclic structure and often contain d-amino acids. Certain species of fungi produce peptides highly toxic to humans. Among them are α-amanitin, a cyclic peptide that selectively inhibits RNA polymerase II.

Table 3.2. Some peptide hormones.

Name	Number of amino acids	Function
Angiotensin II	8	Hypertensor
Vasopressin	9	Water balance
Oxytocin	9	Uterine contraction
Bradikinin	9	Hypotensor
Kalllidin	10	Hypotensor
Gastrin I	17	Gastric secretion of HCl
Melanocyte stimulant (β MSH)	18	Increases melanin
Secretin	27	Pancreatic juice secretion
Glucagon	29	Hyperglycemic
Calcitonin	32	Reduces Ca level in blood
Cholecystokinin-pancreoenzymin	33	Contraction of gall bladder; secretion of pancreatic enzymes
Adrenocorticotrophin	39	Stimulate adrenal cortex

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The peptide bond is formed by nucleophilic attack on the ester carbonyl group of peptidyl-tRNA (pept-tRNA) bound to the P-site by the α-amino group of aa-tRNA in the A-site of the PT center (Figure 1). The first step is the deprotonation of the α-NH3+ group to create the nucleophilic NH2 group (step 1, Figure 1). The pKa of this group in aa-tRNA is estimated to be around 8 and it is likely that the proton is accepted by water. Subsequent nucleophilic attack of the α-NH2 group on the electrophilic carbonyl group (step 2, Figure 1) leads to the formation of the initial protonated tetrahedral reaction product, T±, which, by deprotonation (step 3), forms the tetrahedral intermediate, T−. Steps 2 and 3 can also take place synchronously, as has been suggested for the PT reaction [6••], that is, T± is not a necessary reaction intermediate. Breakdown of T− (step 4) is initiated by donating a proton back to the leaving oxygen to form the products: P-site deacylated tRNA and A-site pept-tRNA. Catalysis of the PT reaction could involve any of the following mechanisms: proper positioning of the peptidyl and aminoacyl ends of the tRNAs in the active site; general acid-base catalysis during deprotonation (step 3, Figure 1) and protonation (step 4); electrostatic stabilization of the transition state(s) leading to T± or T−.

Figure 1. The chemistry of peptide bond formation. The 3′ ends of P-site pept-tRNA (left) and A-site aa-tRNA (right) are depicted. Step 1: deprotonation of the NH3+ group. Step 2: nucleophilic attack of the NH2 group on the ester carbonyl group and formation of a zwitterionic tetrahedral intermediate, T± [33]. Step 3: deprotonation and formation of the negatively charged tetrahedral intermediate, T−. Step 4: product formation by breakdown of the tetrahedral intermediate upon protonation of the leaving oxygen. Steps 2 and 3 can also take place synchronously. Products are P-site deacylated tRNA (left) and A-site pept-tRNA (right). R1 and R2, amino acid sidechains. Red and blue, groups to be deprotonated or protonated, respectively.

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Peptide bond

Fig. 3.7. Peptide bond.

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Peptides

Peptides consist of a group of amino acids linked through an amide (peptide) bond and can be 2–40 in length. The chemistry of amino acid side chains provides aromatic, polar, nonpolar and charged groups, while the amide (HNCO) and carbonyl (CO) groups along the peptide backbone allow the formation of hydrogen bonds. Peptides can be synthesized chemically by solid phase methods and purified by chromatography techniques allowing their large scale production with required grade for medical applications.

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4.3 Primary structure

4.3.1 Peptide bond

Peptide bonds have a planar trans configuration and undergo very little rotation or twisting around the amide bond that links the α-amino nitrogen of one amino acid to the carbonyl carbon of the next amino acid (Fig. 4.2). This effect is due to amido–imido tautomerization. The partial double-bond character of the N–C bond in the transition state probably best represents what exists in nature. Electrons are shared by the nitrogen and oxygen atoms, and the N–C and C–O bonds are both (roughly) “one-and-one-half” bonds (intermediate between single and double). The short carbonyl carbon–nitrogen bond length, 0.132 nm (the usual carbon–nitrogen single bond length is 0.147 nm), is consistent with the partial double-bond character of the peptide linkage. The planarity and rigidity of the peptide bond are accounted for by the fact that free rotation cannot occur around double bonds.

Figure 4.2. Geometry of a peptide (amide) linkage.

Whereas most peptide bonds exist in the trans configuration to keep the side chains (R-groups) as far apart as possible, the peptide bond that involves the –NH group of the rigid pyrrolidone ring of proline can occur in both trans and cis arrangements (Fig. 4.3). However, X-ray data suggest that the trans form occurs more frequently in proteins than does the cis form. It has been further postulated that some proline residues (known as “permissive” proline residues) can exist in either the cis or trans configuration. For example, of the four proline residues of ribonuclease A, at least two are thought to be in the trans configuration in order to form a native structure, whereas one or both of the other residues may be accommodated in either the cis or trans configuration.

Figure 4.3. The trans and cis configurations of peptide bonds involving proline.

The bonds on either side of the α-carbon (i.e., between the α-carbon and the nitrogen, and between the α-carbon and the carbonyl carbon) are strictly single bonds. Rotation is possible around these single bonds. The designation for the angle of rotation of the α-carbon–nitrogen bond is ϕ, whereas that of the α-carbon–carbonyl carbon bond is ψ. Although theoretically an infinite number of ϕ and ψ angles are possible around single bonds, only a limited number of ϕ and ψ angles are actually possible in proteins. A polypeptide has specific ϕ and ψ values for each residue that determines its conformation.

4.3.2 Determination of primary structure

The primary structure of a protein can be determined through the following steps (Fig. 4.4):

Figure 4.4. Illustration of steps involved in the determination of the amino acid sequence of a protein. Instead of using Edman degradation to determine amino acid sequences, Tandem mass spectrophotometry analysis can be employed. Ionized peptide fragments generated by collisions with inert gas atoms in a collision cell are separated by a mass difference of one amino acid due to the fragmentation occurring at the amide bonds of the peptide. By comparing the mass to charge ratio (m/z) data of the peptide fragments with already known data of specific amino acid, the peptide sequence can be deduced.

Obtain a pure sample of the protein of interest. Proteins can be separated on the basis of their size by dialysis, gel filtration, and membrane filtration. Proteins can also be separated by chromatographic techniques such as gas and liquid chromatography, which are based on the differential partitioning of solute molecules due to differences in their affinity for a moving solvent phase and a fixed or supportive phase. Liquid chromatography includes ion exchange and affinity chromatography. Electrophoresis can separate charged proteins on the basis of their different mobilities in an electric field. In particular, the capillary electrophoresis technique has been used to analyze and separate proteins that differ only slightly in amino acid composition or glycosylation.

Determine the amino acid composition and molecular weight of the pure protein. From amino acid composition and molecular weight data, calculate the number of residues of each amino acid present per protein molecule to the nearest whole number. To determine the amino acid composition, the protein is completely hydrolyzed by acid (6N HCl, 24 hours or longer at 110°C, under vacuum or inert gas) to its constituent amino acids. The protein hydrolysate is then separated by cation exchange chromatography on a Dowex 50 resin, which binds all the protonated amino acids. The amino acids are differentially eluted by the use of a pH gradient varying between pH 3 and 5. As each amino acid elutes, it is reacted with ninhydrin to produce a color for spectroscopic detection or o-phthalaldehyde for fluorescence detection.

Fragmentation of the polypeptide by enzymatic or chemical hydrolysis and separation of the peptide mixtures into individual fragments. Hydrolysis of a protein into peptides can be accomplished by group-specific chemical and enzymatic reagents (Table 4.1).

Table 4.1. Hydrolysis of polypeptides at specific sites.


Cleavage reagent	Cleavage site
Enzymes	AA1	AA2
Trypsin	Lys, Arg
Chymotrypsin	Phe, Tyr, Trp
Thermolysin		Leu, IIe, Val
Thrombin		Arg
Elastase	Ala, Gly, Ser, Val	Pro
Pepsin	Leu, Phe, Tyr, Trp	(not Pro)
Chemicals
Cyanogen bromide	Met
2-Nitro-5-thiocyanobenzoate		Cys
N-Bromosuccinimide	Tyr, Trp
Hydroxylamine	Asn	Gly
O-Iodosobenzoate		Trp
Mild acid (70% HCOOH or 0.1N HCI)	Asp	Pro

Sequence each fragment by sequential Edman degradation. Determination of the N-terminal residue is carried out by labeling the free unprotonated α-amino group with the Edman reagent (phenylisothiocyanate, PITC). PITC reacts under basic conditions with the free α-amino group to form a phenylthiocarbamoyl peptide (Fig. 4.5). Treatment with anhydrous acid yields the labeled terminal amino acid residue plus the remainder of the peptide. In this process, the terminal amino acid is cyclized to the corresponding phenylthiohydantoin derivative (PTH-amino acid), which can be identified by chromatographic procedures such as reverse-phase high-pressure liquid chromatography. After the initial Edman degradation, the remaining peptide group is available for another cycle of the procedure. This procedure can thus be used in a stepwise manner to establish the sequence of amino acids in a peptide starting from the N-terminal (Fig. 4.6).

Figure 4.5. Labeling of the N-terminal residue by the Edman degradation.

Figure 4.6. Determination of N-terminal residue of a polypeptide by the Edman procedure. After removal of the N-terminal amino acid, the remainder of the peptide remains intact and a new N-terminal amino acid is available for removal by the next reaction cycle.

Analysis of overlapping peptide sequences and assembly of the original protein sequence. Once the sequence of digested peptides has been determined by the sequential Edman procedure, the proper arrangement of individual peptides in the protein can be established by identifying overlapping sequences between peptides obtained by different cleavage procedures (Fig. 4.7).

Figure 4.7. Deduction of peptide sequence from analysis of overlapping sequences of component peptides.

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1.1 Protein Structure and Reactivity

Amino Acids

Peptides and proteins are composed of amino acids polymerized together through the formation of peptide (amide) bonds. The peptide-bonded polymer that forms the backbone of polypeptide structure is called the α-chain. The peptide bonds of the α-chain are rigid planar units formed by the reaction of the α-amino group of one amino acid with the α-carboxyl group of another (Figure 2.1). The peptide bond possesses no rotational freedom due to the partial double-bond character of the carbonyl–amino amide bond. The bonds around the α-carbon atom, however, are true single bonds with considerable freedom of movement.

Figure 2.1. Rigid peptide bonds link amino acid residues together to form proteins. Other bonds within the polypeptide structure may exhibit considerable freedom of rotation.

The sequence and properties of the amino acid constituents determine protein structure, reactivity, and function. Each amino acid is composed of an amino group and a carboxyl group bound to a central carbon, termed the α-carbon. Also bound to the α-carbon are a hydrogen atom and a side chain unique to each amino acid (Figure 2.2). There are 20 common amino acids found throughout nature, each containing an identifying side chain of particular chemical structure, charge, hydrogen bonding capability, hydrophilicity (or hydrophobicity), and reactivity. The side chains do not participate in polypeptide formation and are thus free to interact and react with their environment.

Figure 2.2. Individual amino acids consist of a primary (α) amine, a carboxylic acid group, and a unique side-chain structure (R). At physiological pH, the amine is protonated and bears a positive charge, while the carboxylate is ionized and possesses a negative charge.

Amino acids may be grouped by type depending on the characteristics of their side chains. There are seven amino acids that contain aliphatic side chains, which are relatively nonpolar and hydrophobic: glycine, alanine, valine, leucine, isoleucine, methionine, and proline (Figure 2.3). Glycine is the simplest amino acid, its side chain consisting of only a hydrogen atom. Alanine is next in line, possessing just a single methyl group for its side chain. Valine, leucine, and isoleucine are slightly more complex with three or four carbon branched-chain constituents. Methionine is unique in that it is the only reactive aliphatic amino acid, containing a thioether group at the terminus of its hydrocarbon chain. Proline is actually the only imino acid. Its side chain forms a pyrrolidine ring structure with its α-amino group. Thus, it is the only amino acid containing a secondary α-amine. Due to its unique structure, proline often causes severe turns in a polypeptide chain. Proteins rich in proline, such as collagen, have tightly formed structures of high density. Collagen also contains a rare derivative of proline, 4-hydroxyproline, found in only a few other proteins. Proline, however, cannot be accommodated in normal α-helical structures, except at the ends where it may create the turning point for the chain. Poly-proline α-helical structures have been formed, but the structural characteristics of these artificial polypeptides are quite different from native protein helices.

Figure 2.3. Common aliphatic amino acids.

Phenylalanine and tryptophan contain aromatic side chains that, like the aliphatic amino acids, are also relatively nonpolar and hydrophobic (Figure 2.4). Phenylalanine is unreactive toward common derivatizing reagents, whereas the indolyl ring of tryptophan is quite reactive, if accessible. The presence of tryptophan in a protein contributes more to its total absorption at 275 to 280 nm on a mole-per-mole basis than any other amino acid. The phenylalanine content, however, adds very little to the overall absorbance in this range.

Figure 2.4. The two aromatic amino acids that have nonpolar and nonionizable side-chain groups.

All of the aliphatic and aromatic hydrophobic residues are often located at the interior of protein molecules or in areas that interact with other nonpolar structures such as lipids. They usually form the hydrophobic core of proteins and are not readily accessible to water or other hydrophilic molecules.

There is another group of amino acids that contains relatively polar constituents and are thus hydrophilic in character. Asparagine, glutamine, threonine, and serine (Figure 2.5) are usually found in hydrophilic regions of a protein molecule, especially at or near the surface where they can be hydrated with the surrounding aqueous environment. Asparagine, threonine, and serine are often found post-translationally modified with carbohydrate in N-glycosidic (asp) and O-glycosidic linkages (thr and ser). Though these side chains are enzymatically derivatized in nature, the hydroxyl and amide portions have relatively the same nucleophilicity as that of water and are therefore difficult to modify with common reagent systems under aqueous conditions.

Figure 2.5. The four amino acids with polar, uncharged side chains. The arrows show the attachment points for carbohydrate that may be present in post-translational modifications on glycoproteins.

The most significant amino acids for modification and conjugation purposes are the ones containing ionizable side chains: aspartic acid, glutamic acid, lysine, arginine, cysteine, histidine, and tyrosine (Figure 2.6). In their unprotonated state, each of these side chains can be potent nucleophiles to engage in addition reactions (see the discussion on nucleophilicity below).

Figure 2.6. The amino acids with ionizable side chain groups possess some of the most important functional groups for bioconjugate applications. The C- and N-terminals of each polypeptide chain also are included in this group.

Both aspartic and glutamic acids contain carboxylate groups that have similar ionization properties to the C-terminal α-carboxylate. The theoretical pKa of the β-carboxyl of aspartic acid (3.7–4.0) and the γ-carboxyl of glutamic acid (4.2–4.5) are somewhat higher than the α-carboxyl groups at the C-terminal of a polypeptide chain (2.1–2.4). At pH values above their pKa, these groups are generally ionized to negatively charged carboxylates. Thus, at physiological pH, they contribute to the overall negative charge contribution of an intact protein (see following section).

Carboxylate groups in proteins may be derivatized through the use of amide bond-forming agents or through active ester or reactive carbonyl intermediates (Figure 2.7). The carboxylate actually becomes the acylating agent to the modifying group. Amine-containing nucleophiles can couple to an activated carboxylate to give amide derivatives. Hydrazide compounds react similarly to amines. While a thiol group is reactive toward an activated carboxylate and results in a thioester linkage, it forms relatively unstable derivatives, which can exchange with other nucleophiles such as amines or hydrolyze in aqueous solutions.

Figure 2.7. Derivatives of carboxylic acids can be prepared through the use of active intermediates that react with target functional groups to give acylated products.

Lysine, arginine, and histidine have ionizable amine-containing side chains that, along with the N-terminal α-amine, contribute to a protein’s overall net positive charge. Lysine contains a straight four-carbon chain terminating in a primary amine group. The ε-amine of lysine differs in pK1 from the primary α-amines by having a slightly higher ionization point (pKa of 9.3–9.5 for lysine versus pKa of 7.6–8.0 for α-amines). At pH values lower than the pKa of these groups, the amines are generally protonated and possess a positive charge. At pH values greater than the pKa, the amines are unprotonated and contribute no net charge. Arginine contains a strongly basic chemical constituent on its side chain called a guanidino group. The ionization point of this residue is so high (pKa>12.0) that it is virtually always protonated and carriers a positive charge. The histidine side chain is an imidazole ring that is potentially protonated at slightly acidic pH values (pKa=6.7–7.1). Thus, at physiological pH, these residues contribute to the overall net positive charge of an intact protein molecule.

The amine-containing side chains in lysine, arginine, and histidine typically are exposed on the surface of proteins and can be derivatized with ease. The most important reactions that can occur with these residues are alkylation and acylation (Figure 2.8). In alkylation, an active alkyl group is transferred to the amine nucleophile with loss of one hydrogen. In acylation, an active carbonyl group undergoes addition to the amine. Alkylating reagents are highly varied and the reaction with an amine nucleophile is difficult to generalize. Acylating reagents, however, usually proceed through a carbonyl addition mechanism as shown in Figure 2.9. The imidazole ring of histidine is also an important reactive species in electrophilic reactions, such as in iodination using radioactive 125I or 131I (Chapter 12, Section 2).

Figure 2.8. Derivatives of amines can be prepared from acylating or alkylating agents to give amide, secondary amine, or tertiary amine bonds.

Figure 2.9. The mechanism of acylation proceeds through the attack of a nucleophile, generating a tetrahedral intermediate, which then goes on to form the product.

Cysteine is the only amino acid containing a sulfhydryl group. At physiological pH, this residue is normally protonated and possesses no charge. Ionization only occurs at high pH (pKa=8.8–9.1) and results in a negatively charged thiolate residue. The most important reaction of cysteine groups in proteins is the formation of disulfide crosslinks with another cysteine molecule. Cysteine disulfides (called cystine residues) are often key points in stabilizing protein structure and conformation. They frequently occur between polypeptide subunits, creating a covalent linkage to hold two chains together. Cysteine and cystine groups are relatively hydrophobic and usually can be found within the core of a protein. For this reason, it is often difficult to fully reduce the disulfides of large proteins without a deforming agent present to open up the inner structure and make them accessible (see Section 4.1).

Cysteine sulfhydryls and cystine disulfides may undergo a variety of reactions, including alkylation to form stable thioether derivatives, acylation to form relatively unstable thioesters, and a number of oxidation and reduction processes (Figure 2.10). Derivatization of the side chain sulfhydryl of cysteine is one of the most important reactions of modification and conjugation techniques for proteins.

Figure 2.10. Sulfhydryl groups may undergo a number of additional reactions, including acylation and alkylation. Thiols also may participate in redox reactions, which generate reversible disulfide linkages.

Tyrosine contains a phenolic side chain with a pKa of about 9.7 to 10.1. Due to its aromatic character, tyrosine is second only to tryptophan in contributing to a protein’s overall absorptivity at 275 to 280 nm. Although the amino acid is only sparingly soluble in water, the ionizable nature of the phenolic group makes it often appear in hydrophilic regions of a protein—usually at or near the surface. Thus, tyrosine derivatization proceeds without much need for deforming agents to further open protein structure.

Tyrosine may be targeted specifically for modification through its phenolate anion by acylation, through electrophilic reactions such as the addition of iodine or diazonium ions, and by Mannich condensation reactions. The electrophilic substitution reactions on tyrosine’s ring all occur at the ortho position to the –OH group (Figure 2.11). Most of these reactions proceed effectively only when tyrosine’s ring is ionized to the phenolate anion form.

Figure 2.11. Tyrosine residues are subject to nucleophilic and electrophilic reactions. The unprotonated phenolate ion may be alkylated or acylated using a variety of bioconjugate reagents. Its aromatic ring also may undergo electrophilic addition using diazonium chemistry or Mannich condensation, or be halogenated with radioactive isotopes such as 125I.

In summary, protein molecules may contain up to nine amino acids that are readily derivatizable at their side chains: aspartic acid, glutamic acid, lysine, arginine, cysteine, histidine, tyrosine, methionine, and tryptophan. These nine residues contain eight principal functionalities with sufficient reactivity for modification reactions: primary amines, carboxylates, sulfhydryls (or disulfides), thioethers, imidazoles, guanidinyl groups, and phenolic and indolyl rings. All of these side-chain functionalities in addition to the N-terminal α-amino and the C-terminal α-carboxylate form the full complement of polypeptide reactivity within proteins (Figure 2.12).

Figure 2.12. The more important polypeptide functional groups are represented by these nine amino acids. Bioconjugate chemistry may occur through the C- and N-terminals of each polypeptide chain, the carboxylate groups of aspartic and glutamic acids, the ε-amine of lysine, the guanidino group of arginine, the sulfhydryl group of cysteine, the phenolate ring of tyrosine, the indol ring of tryptophan, the thioether of methionine, and the imidazole ring of histidine.

Nucleophilic Reactions and the pI of Amino Acid Side Chains

Ionizable groups within proteins can exist in one of two forms: protonated or unprotonated. Carboxylate groups below their pKa values exist in the protonated state and are therefore in the conjugate acid form and carry no charge. However, at pH values above the pKa of the carboxylic group, the acid is ionized and therefore unprotonated to a negative charge. This same relationship is true of the –OH group on the phenol ring of tyrosine. At pH values below its pKa, tyrosine’s side chain is uncharged. Above the pKa, however, the hydrogen ionizes off, leaving a negatively charged phenolate. Conversely, amine nucleophiles below their pKa values are in a protonated state and possess a positive charge. At pH values above the pKa of the amino group, it is then ionized and unprotonated to neutrality.

Each type of ionizable group in proteins will have a unique pKa based upon the theoretical value for the amino acid and modulated from that value by its own surrounding microenvironment. Minute environmental changes will cause amine-containing residues at different structural locations to have different ionization potentials, even if the groups are otherwise chemically identical.

Thus, the actual pKa of each ionizable group within protein molecules may range considerably lower or higher than the theoretical values as the microenvironment of individual groups changes. Identical side chains in differing parts of a protein molecule may have widely varying pKa values depending on the immediate chemical milieu. Such factors as the presence of other amino acid side chains in the vicinity, salts, buffers, temperature, ionic strength, and other effects of the solvent medium all play crucial roles in creating microenvironmental changes that affect the ionization potential of these groups (Tanford and Hauenstein, 1956; Schewale and Brew, 1982).

The Henderson–Hasselbalch equation (1) explains the relationship of pH and pKa to the relative ratios of protonated (acid) and unprotonated (base) forms of an ionizable group. Note that the ionized form of such a group does not have to possess a negative charge, as in the case of unprotonated primary amines. Indeed, in that instance it is the protonated amine that bears a charge of positive one. According to the mathematical implications of this equation, an ionizable group at its pKa value is exactly 50% ionized. This means that aspartic acid side chains placed in a medium with a pH equal to its pKa should have half of its carboxylates ionized to a negative charge and half of them unionized with no charge.

(1)pH=pKa+log{[base]/[acid]}

Further implications of this equation are that at one pH unit below or above the pKa, an ionizable group will be 91% unionized (protonated) or 91% ionized (unprotonated), respectively. Two pH units below or above translate to a 99% unionized or 99% ionized state.

The absolute ratio of protonated : unprotonated forms will change from this theoretical approach based upon the microenvironment each group experiences. The reactivity of amino acid side chains is directly related to them being in an unprotonated or ionized state. Many reactions of modification and conjugation occur efficiently only when the nucleophilic species is in an ionized form. As the unprotonated form increases in concentration, the relative nucleophilicity of the ionizable group increases. Many of the reactive groups commonly used for protein modification will couple in greater yield as the pH of the reaction is raised closer to the pKa of the ionizable target. However, continuing to increase the pH beyond the pKa may not be necessary for increased yield, and may even be detrimental, because many reactive groups will begin to lose activity through hydrolysis at high pHs.

A nucleophile is any atom containing an unshared pair of electrons or an excess of electrons able to participate in covalent bond formation. Nucleophilic attack at an atomic center of electron deficiency or positive charge is the basis for many of the coupling reactions that occur in chemical modification. Thus, an uncharged amine group is a more powerful nucleophile than the protonated form bearing a positive charge. Likewise, a negatively charged carboxylate has greater nucleophilicity than its uncharged, protonated conjugate acid form. In addition, an unprotonated thiolate, bearing a negative charge (RS−), is a much more powerful nucleophile than its protonated, uncharged sulfhydryl form.

According to the theory of nucleophilicity (Edwards and Pearson, 1962; Bunnett, 1963; Pearson et al., 1968), the relative order of nucleophilicity relative to the major groups in biological molecules can be summarized as follows:

R-S−>R-SH

R-NH2>R-NH3+

R-COO−>R-COOH

R-O−>R-OH

R-OH=H-OH

and finally,

R-S−>R-NH2>R-COO−=R-O−

Using these relationships, it is obvious that the strongest nucleophile in protein molecules is the sulfhydryl group of cysteine, particularly in the ionized, thiolate form. Next in line are the amine groups in their uncharged, unprotonated forms, including the α-amines at the N-terminals, the ε-amines of lysine side chains, the secondary amines of histidine imidazolyl groups and tryptophan indole rings, and the guanidino amines of arginine residues. Finally, the least potent nucleophiles are the oxygen-containing ionizable groups including the α-carboxylate at the C-terminal, the β-carboxyl of aspartic acid, the γ-carboxyl of glutamic acid, and the phenolate of tyrosine residues.

According to the theoretical pKa values for the ionizable side chains of amino acids, nucleophilic substitution reactions involving primary amines or sulfhydryl groups on proteins should not be efficient below a pH of about 8.5 (Table 2.1). In practice, however, reactions can be carried out with these groups in high yield at pH values not much higher than neutrality. This discrepancy relates to the changes in pKa due to microenvironmental effects experienced by the residues within the three-dimensional structure of the protein molecule. In reality, the ε-amine groups on lysine side chains within proteins, having theoretical pKas of over 10, nonetheless exist in sufficient quantity in an unprotonated form even at a pH of 7.2 that modification easily occurs.

Table 2.1. pKa of Ionizable Amino Acids

Group Location	Functionality	pKa Range
α-Amine; N-terminus		7.6–8
Lysine’s ε-amine		9.3–9.5
Histidine’s imidazolyl nitrogen		6.7–7.1
Arginine’s guanidinyl group		>12
Tyrosine’s phenolic hydroxyl		9.7–10.1
α-Carboxyl; C-terminus		2.1–2.4
Aspartic acid’s γ-carboxyl		3.7–4
Gutamic acid’s γ-carboxyl		4.2–4.5
Cysteine’s sulfhydryl		8.8–9.1

One important point should be noted, however. The changes that occur in the pKa of ionizable groups in protein molecules due to microenvironmental effects sometimes make it difficult to select certain residues for modification simply by careful modulation of reaction pH. For instance, at least in theory, overlap of the pKa range for sulfhydryls and amine-containing residues would eliminate any chance of directing a reaction toward –SH groups solely by adjusting the pH of the reaction medium. However, because of the microenvironmental changes that occur in complex biomolecules, pH sometimes can be used along with the right reactive group to target thiols without amine modification. Thus, in practice, to effectively site-direct a modification reaction, the proper choice of reactive group and reaction conditions can result in highly discrete conjugation to certain sites within proteins.

Secondary, Tertiary, and Quaternary Structure

Amino acids are linked through peptide bonds to form long polypeptide chains. The primary structure of protein molecules is simply the linear sequence of each residue along the α-chain. Each amino acid in the chain interacts with surrounding groups through various weak, noncovalent interactions and through its unique side-chain functionalities. Noncovalent forces such as hydrogen bonding and ionic and hydrophobic interactions combine to create each protein’s unique organization.

The sequence and types of amino acids and the way that they are folded provide protein molecules with specific structure, activity, and function. Ionic charge, hydrogen bonding capability, and hydrophobicity are the major determinants for the resultant three-dimensional structure of protein molecules. The α-chain is twisted, folded, and formed into globular structures, α-helicies, and β-sheets based upon the side-chain amino acid sequence and weak intramolecular interactions such as hydrogen bonding between different parts of the peptide backbone (Figure 2.13). Major secondary structures of proteins such as α-helicies and β-sheets are held together solely by massive hydrogen bonding created through the carbonyl oxygens of peptide bonds interacting with the hydrogen atoms of other peptide bonds (Figure 2.14).

Figure 2.13. The α-chain structure of alkaline phosphatase illustrates the complex nature of polypeptide structure within proteins (Kim and Wyckoff, 1991).

Figure 2.14. Secondary structures within proteins may be stabilized through hydrogen bonding between adjacent α-chains, forming β-sheet conformations.

In addition, negatively charged residues may become bonded to positively charged groups through ionic interactions. Nonpolar side chains may attract other nonpolar residues and form regions of hydrophobicity to the exclusion of water and other ionic groups. Occasionally, disulfide bonds are also found holding different regions of the polypeptide chain together. All of these forces combine to create the secondary structure of proteins, which is the way the polypeptide chain folds in local areas to form larger, sometimes periodic structures.

On a larger scale, the unique folding and structure of one complete polypeptide chain is termed the tertiary structure of protein molecules. The difference between local secondary structure and complete polypeptide tertiary structure is arbitrary and sometimes of little practical difference.

Larger proteins often contain more than one polypeptide chain. These multi-subunit proteins have a more complex shape, but are still formed from the same forces that twist and fold the local polypeptide. The unique 3-dimensional interaction between different polypeptides in multi-subunit proteins is called the quaternary structure. Subunits may be held together by noncovalent contacts, such as hydrophobic or ionic interactions, or by covalent disulfide bonds formed from the cysteine residue of one polypeptide chain being crosslinked to a cysteine sulfhydryl of another chain (Figure 2.15).

Figure 2.15. Polypeptide chains may be bound together through disulfide linkages occurring between cysteine residues within each subunit.

Thus, aside from the covalently polymerized α-chain itself, the majority of protein structure is determined by weaker, non-covalent interactions that potentially can be disturbed by environmental changes. It is for this reason that protein structure can be easily disrupted or denatured by fluctuations in pH or temperature or by substances that can alter the structure of water, such as detergents or chaotropes.

Not surprisingly, chemical modification to the amino acid constituents of a polypeptide chain may also cause significant disruption in the overall three-dimensional structure of a protein. If amino acid residues critical to folding near functionally important regions are modified with chemical groups that change the charge, hydrophilicity, or hydrogen bonding character of the polypeptide chain, protein structure may be altered and activity may be compromised. This concept will be discussed further in subsequent sections.

Prosthetic Groups, Cofactors, and Post-Translational Modifications

Proteins may contain structures other than polypeptide chains that are important for biological function. Prosthetic groups and cofactors are small organic compounds that are sometimes tightly bound to a protein and aid in forming the active center. A prosthetic group is usually carried within the three-dimensional protein structure in a firm-fitting pocket or even attached through covalent bonds, such as the heme ring associated with cytochrome c molecules which is bonded through thioether linkages with adjacent cysteine residues (Figure 2.16). Cofactors, by contrast, may be bound only transiently to proteins during periods of activity. Enzymes often require cofactors to act as donors or acceptors of chemical groups that are added to or cleaved from a substrate molecule. Some common cofactors are ATP, ascorbic acid, coenzyme A, NAD, NADP, FAD, FMN, and biotin. Sometimes, the enzyme cofactor is also an energy source for the catalytic reaction, as in the case of ATP dependent reactions.

Figure 2.16. The heme ring of cytochrome c is a non-amino-acid, prosthetic group bound to the protein through two cysteine residues.

Frequently, metal ions are associated with the prosthetic group or cofactor. Heme rings usually contain a chelated iron atom. Occasionally, however, these metals are merely bound within folded polypeptide regions with no additional organic constituents required. Many metal ions are known to participate in enzymatic activity. One or more of the ions of Na, K, Ca, Zn, Cu, Mg, and Mn, as well as Co and Mo, are often required by enzymes to maintain activity.

Prosthetic groups and cofactors, whether organic or metallic, may be removed from a protein to create an inactive apo protein or enzyme. Loss of these groups may occur through environmental changes, such as removing metal ions from solution or adding denaturants to unfold protein structure. In many cases, simply re-introducing the needed group into the surrounding medium can restore full activity.

In addition to small organic molecules or metal ions, proteins may have other components tightly associated with them. Nucleoproteins, for instance, contain noncovalently bound DNA or RNA, as in some of the structural proteins of viruses. Lipoproteins contain associated lipids or fatty acids and may also carry cholesterol, as in the high-density and low-density lipoproteins in serum.

During modification or conjugation reactions, prosthetic groups and other associated molecules may be lost or damaged. Metal ions temporarily may be removed by the inclusion of a chelating agent added to maintain sulfhydryl stability during coupling through the –SH groups of a protein. To restore activity after conjugation, it is necessary to remove the chelator and add the required metal salts. Other changes to the prosthetic carriers may not be so easily corrected. For instance, heme-containing molecules are sensitive to the presence of agents that can form a coordination complex with or modify the oxidation state of the chelated metal ion. Some reagent systems may permanently inactivate the heme-containing protein.

Thus, loss of activity can occur not only through changes to the amino acid constituents of a protein, but through prosthetic group or cofactor loss or damage as well. Most of these potential difficulties can be overcome through careful selection of the reaction conditions and through knowledge of the cofactor dependencies that are critical to the activity of the protein being modified.

Post-translational modifications to protein structure are covalent changes that occur as the result of controlled enzymatic reactions or due to chemical reactions not under enzymatic regulation. One of the most common cellular modifications performed on proteins after ribosomal synthesis is glycosylation. Proteins newly synthesized on ribosomes may be transported to the Golgi apparatus, where specific glycosyl transferases catalyze the coupling of carbohydrate residues to the polypeptide chains. Glycoproteins and mucoproteins are formed by the coupling of polysaccharides through O-glycosidic linkages to serine, threonine, or hydroxylysine and through N-glycosidic linkages with the amide side-chain group of asparagine.

The structure of most glycoprotein carbohydrate is branched with the sugars mannose, N-acetylglucosamine (GlcNAc), sialic acid, glactose, and l-fucose being prevalent. Asparagine-linked polysaccharides are well characterized and are known to be constructed of a core unit consisting of three mannose residues and two N-acetylglucosamine residues. The GlcNAc residues are bound to the Asp side-chain amide nitrogen through a β1 linkage (Kornfield and Kornfield, 1985). The three mannose groups then usually form the first branch point in the oligosaccharide chain (Section 2.1.2).

The content by weight of carbohydrate in glycoproteins may vary from only a few percent to over 50% in some proteins in mucous secretions. Although the function of the polysaccharide in most glycoproteins is unknown, in some cases it may provide hydrophilicity, recognition, and points of non-covalent interaction with other proteins through lectin-like affinity binding.

The presence of carbohydrate on protein or peptide molecules can provide important points of attachment for modification or conjugation reactions. Coupling exclusively through polysaccharide chains can often direct the reaction away from active centers or critical points in the polypeptide chain, thus preserving activity. Polysaccharides can be specifically targeted on glycoproteins through mild sodium periodate oxidation. Periodate cleaves adjacent hydroxyl groups in sugar residues to create highly reactive aldehyde functionalities (Chapter 3, Section 4.4). The level of periodate addition can be adjusted to selectively cleave only certain sugars in the polysaccharide chain. For instance, a concentration of 1-mM sodium periodate at temperatures less than 4°C specifically oxidizes sialic acid residues to contain aldehydes, leaving all other monosaccharides untouched. Increasing the concentration to 10-mM and carrying out the reaction at room temperature, however, will cause oxidation of other sugars in the carbohydrate chain, including galactose and mannose. The generated aldehydes can then be used in coupling reactions with amine-or hydrazide-containing molecules to form covalent linkages. Amines can react with formyl groups under reductive amination conditions using a suitable reducing agent such as sodium cyanoborohydride. The result of this reaction is a stable secondary amine linkage (Chapter 3, Section 5.3). Alternatively, hydrazides spontaneously react with aldehydes to form hydrazone linkages, although the addition of a reducing agent increases the efficiency of the reaction (Chapter 3, Section 5.1).

Another form of post-translational modification that may add carbohydrate to a polypeptide is non-enzymatic glycation. This reaction occurs between the reducing ends of sugar molecules and the amino groups of proteins and peptides. See Section 2.1 in this Chapter for further details and the reaction sequence behind this modification.

Protecting the Native Conformation and Activity of Proteins

The goal of most protein modification or conjugation procedures is to create a stable product with good retention of the native state and activity. Ideally, any derivatization should result in a protein that performs exactly as it would in its unmodified form, but with the added functionality imparted by whatever is conjugated to it. Thus, an antibody molecule tagged with a fluorophore should retain its ability to bind to antigen and also have the added functionality of fluorescence.

One of the best ways to ensure retention of activity in protein molecules is to avoid carrying out chemistry at the active center. The active center is that portion of the protein where ligand, antigen, or substrate binding occurs. In simpler terms, the active center (or active site) is that part that has specific interaction with another substance (Means and Feeney, 1971). For the preparation of enzyme derivatives, it is important to protect the site of catalysis where conversion of substrate to product happens. For instance, when working with antibody molecules, it is crucial to stay away from the two antigen binding sites.

The best chemical procedures avoid the active site by selecting functional groups away from that area or by protecting the site through the incorporation of additives. In some cases, the inclusion of substrates, cofactors, ligands, inhibitors, or antigens in the modification reaction will protect the active site. Addition of the appropriate substance can bind the active site and mask it from modification by crosslinking agents. In enzyme derivatization procedures, this is often just a matter of adding a reversible inhibitor or substrate analog. For instance, when working with alkaline phosphatase merely carrying out the reaction in phosphate buffer protects the active center from chemical modification, since phosphate ions bind in the catalytic site. With trypsin, the incorporation of benzamidine similarly masks and protects the active site.

However, protecting the antigen binding sites on an antibody molecule by using this method is often more difficult. Inclusion of antigen to mask the binding sites is effective in blocking these areas, but it may also cause irreversible crosslinking of the antigen to the antibody. This is especially true when the antigen is a peptide or a protein having the same chemical functionalities as the antibody. Any modification reactions that are directed at the antibody may modify the antigen as well. Therefore, only use this method if the antigen is lacking in the chemical targets that are going to be used on the antibody. For instance, if the polysaccharide chains on the antibody are targeted for modification, then using a protein antigen that does not contain carbohydrate to block the antigen binding sites may work well.

An equally effective method of protecting the activity of a protein is by using site-directed reactions that result in modifications away from the active center. In some cases, specific functionalities are known to be present only at restricted sites within the three-dimensional structure of a protein. If these functionalities are not present close to the active site, then using them exclusively for modification reactions should ensure good retention of activity. For instance, sulfhydryl groups or carbohydrate chains are often present in limited quantity and in specific regions on a protein. Selecting reagent systems that target these groups ensures derivatization only at restricted sites within the protein molecule, thus potentially avoiding the active center.

Surprisingly, the goal of some protein crosslinking schemes is to somewhat alter the native presentation of the conjugate. This is especially true in hapten–carrier conjugation as used for immunogen or vaccine preparation. In this case, the main objective is to modify the environment of the hapten to create an immunological response in vivo. A hapten is usually a small molecule that is not able to generate an immune response on its own, but can react with the products of such a response once generated. Most often these products are antibodies having binding specificity for the hapten.

The complexities involved in achieving a successful conjugation strategy are best illustrated in the problems and concerns dealing with hapten–carrier conjugation. In order to produce the initial immune response to a small molecule, the hapten is typically coupled to a larger protein that can generate a response on its own. In simple terms, the larger carrier protein confers immunogenicity to the smaller hapten. The native presentation of the hapten is altered toward the immune system, thus creating the immune response.

The site of attachment of the hapten to the carrier and the nature of the crosslinker are both important to the specificity of the resultant antibodies generated against it. For proper recognition, the hapten must be coupled to the carrier with the appropriate orientation. For an antibody subsequently to recognize the free hapten without the attached carrier, the hapten–carrier conjugate must present the hapten in an exposed and accessible form. Optimal orientation is often achieved by directing the crosslinking reaction to specific sites on the hapten molecule. With peptide haptens, this is typically carried out by attaching a terminal cysteine residue during synthesis. This provides a free thiol group on one end of the peptide for conjugation to the carrier. Crosslinking through this group provides hapten attachment only at one end, therefore ensuring consistent orientation.

In hapten–carrier conjugation, the goal is not to maintain the native state or stability of the carrier, but to present the hapten in the best possible way to the immune system. In reaching this goal, the choice of conjugation chemistry may control the resultant titer, affinity, and specificity of the antibodies generated against the hapten. It may be important in some cases to choose a crosslinking agent containing a spacer arm long enough to present the antigen in an unrestricted fashion. It may also be important to control the density of the hapten on the surface of the carrier. Too little hapten substitution may result in little or no response. A hapten density that is too high may actually cause immunological suppression and decrease the response. In addition, the crosslinker itself may generate an undesired immune response. Fortunately, for the majority of hapten–carrier conjugation problems, a few main crosslinking techniques provide a workable compromise to solving all these concerns and ultimately generating an effective immune response (Chapter 19).

Oxidation of Amino Acids in Proteins and Peptides

The modification of amino acids in proteins and peptides by oxidative processes plays a major role in the development of disease and in aging (Kim et al., 1985; Tabor and Richardson, 1987; Halliwell and Gutteridge, 1989, 1990Halliwell and Gutteridge, 1989Halliwell and Gutteridge, 1990; Stadtman, 1992). Tissue damage through free-radical oxidation is known to cause various cancers, neurological degenerative conditions, pulmonary problems, inflammation, cardiovascular disease, and a host of other problems. Oxidation of protein structures can alter activity, inhibit normal protein interactions, modify amino acid side chains, cleave peptide bonds, and even cause crosslinks to form between proteins.

Due to their abundance in cells relative to other biological molecules, proteins are one of the primary targets of oxidation in vivo. However, sometimes oxidation reactions involving proteins and peptides are thought of solely as the creation of disulfides from thiols on cysteine residues. This is certainly an important form of oxidation that can affect protein structure and function or even cause problems relevant to bioconjugation reactions. The presence of an accessible free thiol on a protein in an aqueous solution can be highly unstable to rapid oxidation unless precautions are taken to prevent disulfide formation. Dissolved oxygen and other potentially catalytic components, such as certain metal salts, can quickly result in disulfides being formed within a protein or between different protein molecules.

From a broader perspective, protein oxidation can result in covalent modification at many sites other than just at cysteine thiols. The earliest reports on protein oxidation date from the first decade of the 20th century, but it took many more years to characterize these reactions and their products (Dakin, 1906).

The significance of protein oxidation became paramount with the advent of recombinant protein biologics used as human therapeutics. Careful characterization of protein stability is essential to maintaining the efficacy of protein pharmaceuticals. If even a single side-chain amino acid residue becomes oxidized, then a protein therapeutic may not have the same activity in vivo as the unmodified protein.

Oxidation of proteins can result from exposure to oxidative species from many sources: reactive oxygen intermediates caused by metabolic reactions within cells (mitochondrial electron transport function and certain enzymes, such as oxidases, peroxidases, and P-450 enzymes), from the byproducts of oxidative stress reactions in cells (Sayre et al., 2001), or through the presence of strongly oxidizing compounds within a solution—all of these can contribute to selective damage or modification to protein structures. Some examples of chemical agents that can oxidatively modify proteins include hydrogen peroxide (H2O2) and other peroxy compounds, such as perborate and peroxycarbonate; hydroperoxyl radical (HO2·); superoxide anion (O2·−); singlet oxygen (1O2); hydroxyl radical (·OH), periodate (IO4−); metal salts in the presence of oxygen species, such as those of iron (Fe3+ and Fe2+) and copper (Cu2+); ozone (O3); peroxynitrite (ONOO−); hypobromous acid (HOBr); hypochlorous acid (HOCl); performic acid (HC(O)OOH); trichloromethylperoxyl radical (CCl3OO·); under the right conditions, metal-chelating compounds, such as porphyrins, texaphyrins, and FeBABE; and gamma radiation and ultraviolet light. For additional information, see Winterbourn and Kettle, 2000; Baynes and Thorpe, 2000; Greenacre and Ischiropoulos, 2001; Halliwell and Gutteridge, 1989, 1990Halliwell and Gutteridge, 1989Halliwell and Gutteridge, 1990; and Stadtman, 1992.

Singlet oxygen (1O2) differs from the predominant oxygen molecule in that O2 is in the ground state or triplet state and its outer two unshared electrons have parallel spins (sometimes designated 3O2), which is nearly unreactive toward other molecules, while singlet oxygen has increased energy and has its outer electrons transformed into an opposite spin orientation, which is highly reactive. Superoxide (O2·−) is different from singlet oxygen in that it is a reduced form of oxygen having an extra unpaired electron, called a radical. The presence of the radical makes superoxide extremely reactive and highly damaging to proteins and other biological molecules.

Singlet oxygen and superoxide, in addition to their modifying effects on proteins, are also important reactive oxygen species in biological applications, as they are intermediates used in some detection methods and in photodynamic therapy (PDT) for the treatment of cancer. One of the more common compounds used in PDT is Photofrin, which is a mixture of oligomers consisting of ether and ester linkages that combine up to eight porphyrin groups (Misawa et al., 2005). The generation of reactive oxygen species takes place by irradiation with 630-nm wavelength laser light, which also penetrates the skin effectively during therapy. Photoactivation of the Photofrin molecule causes radical initiation to form porphyrin-excited states. Transfer of electrons from the porphyrin groups to molecular oxygen then generates the highly reactive singlet oxygen species. Subsequent radical reactions can also form superoxide and hydroxyl radicals, all of which severely damage tissue in the region of the tumor and ultimately cause cancer cell death.

Another compound used to generate reactive oxygen species for PDT is texaphyrin, which contains a metal-chelating ring structure resembling a porphyrin group (Figure 2.17). Typically, a gadolinium atom is chelated in the texaphyrin center and this complex becomes both a photosensitizing agent and an MRI contrast agent to better visualize tumor locations for irradiation therapy (Donnelly et al., 2004).

Figure 2.17. The texaphyrin–gadolinium chelate structure used as a photosensitizer and MRI contrast agent in the detection and treatment of cancer.

The reactive oxygen species involved with protein oxidation can be generally categorized according to their relative reactivity as follows:

HO⋅,HO2⋅>O2⋅2>ROOH,H2O2>1O2,ClO−,BrO−>O2

Thus, radicals are the most reactive and destructive of protein structure, followed by peroxy derivatives, singlet oxygen, and other oxygen compounds. The oxidative reactivity of some of these oxygen species is so high that just contact of the pure compound with paper or cotton fabrics can cause combustion (e.g., superoxide).

In vitro studies of protein oxidation indicate that virtually all proteins and peptides are susceptible to damage by the radicals ·OH and O2·−. Analysis of protein modification products after oxidation indicates the presence of altered molecular weight (either fragmentation or oligomerization), altered net charge, tryptophan destruction, and the formation of tyrosine dimers (Davies, 1987). Even in the presence of very low concentrations of oxidants (nM), SDS polyacrylamide gel electrophoresis of proteins can indicate multiple bands of higher and lower molecular weight due to oxidative damage.

Transition metals in solution can catalyze the formation of reactive oxygen species that are particularly damaging to proteins and other biomolecules. In a series of reactions, reduced transition metals, such as Fe2+ and Cu1+, can be oxidized by oxygen to produce superoxide and ultimately undergo a Fenton reaction to create hydroxyl radicals (Kim et al., 1985). Transition metal-chelating groups can accelerate this reaction, as demonstrated in the process of hydroxyl radical footprinting of protein interactions using EDTA chelates of iron (see discussion on the reagent FeBABE in Chapter 24, Section 4).

Production of superoxide: Fe2++O2→Fe3++O2⋅−

Production of hydrogen peroxide:2O2⋅−+2H+→H2O2+O2

Production of hydroxyl radical:Fe2++H2O2→Fe3++OH⋅+OH−

The potential sites of oxidation within a protein molecule include the peptide backbone and the side-chain amino acid groups. Hydrogen atom abstraction at the alpha carbon of an amino acid chain can occur upon reaction with an oxidative species to form a radical intermediate. Subsequent reaction can result in peptide bond cleavage and fragmentation of the protein structure, often forming carboxylic acids or carbonyls (aldehydes or ketones). This is the basic mechanism of fragmentation caused by the bifunctional chelating reagent FeBABE. When used in the presence of H2O2 and ascorbic acid, polypeptides will fragment in the neighborhood of interacting proteins.

Amino acid side chains can undergo oxidation through hydrogen abstraction, elimination, or by addition reactions. In the presence of oxygen, aliphatic amino acids usually experience oxidation to a peroxy intermediate, which causes either hydrogen atom abstraction or an elimination reaction resulting in the formation of carbonyls, hydroxyls, or other peroxides (Requena et al., 2001) (Figure 2.18). Aromatic amino acids typically undergo addition reactions following exposure to strong oxidants. An example of this type of reaction is the nitrosation of tyrosine groups in the presence of a peroxynitrite (ONOO−) to create o-nitrotyrosine (see Figure 2.19).

Figure 2.18. Reaction of proline, arginine, and lysine residues with hydroxyl radical results in oxidation of side-chain structures that form carbonyls. Both arginine and proline oxidation will result in the same product being formed.

Figure 2.19. Tyrosine and phenylalanine residues can undergo oxidation to modify their phenyl side chain groups. Tyrosine can form covalent dimers that link two side chains together via a radical reaction. Both tyrosine and phenylalanine can be modified by oxidation to add oxygen-containing groups directly to their aromatic ring.

After exposure to an oxidant, the potential types of oxidation products in proteins and peptides can be extensive (Stadtman and Levine, 2000). Cysteine and methionine undergo a variety of sulfur oxidation reactions to yield cysteine disulfides, methionine sulfoxide, methionine sulfone, and sulfonate products (e.g., cysteic acid) (Ghesquiere et al., 2011) (Figure 2.20). Oxidation with performic acid can be used purposely to convert methionine and cysteine in peptides and proteins to more stable products prior to acid hydrolysis and amino acid analysis. Cysteine and methionine are perhaps the most sensitive amino acids to oxidation, and for this reason they are an early indicator of oxidative damage to proteins.

Figure 2.20. Cysteine and methionine are highly susceptible to oxidation reactions. Cysteine thiols can forms disulfide linkages with other cysteine groups or be oxidized to cysteic acid. Methionine is oxidized very easily to the sulfoxide or sulfone products.

Tyrosine is also easily modified through addition reactions due to the ring activating nature of its phenolic group. Using oxidants, tyrosine’s ring can be chlorinated or iodinated, undergo nitrosation or hydroxylation, and even form tyrosine–tyrosine crosslinks. The last product can be formed purposely by use of a peroxidase in the presence of hydrogen peroxide, and this type of reaction has been studied extensively in the manufacture of phenolic polymer resins (Dordick, 1991). In addition, Fancy et al. (1996) as well as Fancy and Kodadek (1997, 1998)Fancy and Kodadek (1997)Fancy and Kodadek (1998) have applied the oxidation of tyrosine to form dityrosine to the study of protein–protein interactions using nickel-chelated 6xHis-tagged fusion proteins in oxidative environments.

Nitrogen-containing side chains in amino acids can be altered by oxidation forming chloramines or even become deaminated. The result is often the formation of carbonyls (e.g., aldehydes) and hydroxyls. Lee et al. (2006) found that Fe-EDTA-mediated oxidation of human serum albumin resulted in extensive aldehyde and ketone formation from modification of lysine, arginine, histidine, proline, threonine, and aspartic and glutamic acids. Some groups will oxidize and convert to another amino acid altogether. For instance, histidine can be converted to asparagine and proline to hydroxyproline, and tyrosine can be changed to dihydroxyphenylalanine (DOPA) through oxidation reactions.

It is obvious that the oxidation of protein molecules can have detrimental effects on protein structure and function. However, there are some unique methods in bioconjugation wherein controlled and purposeful oxidation is carried out to study protein–protein interactions (Chapter 24, Section 4).

Unfortunately, there are no universal methods to detect all types of protein oxidation, because the products formed can be so diverse in nature. However, some forms of protein oxidation can be assayed using chemical modification (Davies et al., 1999; Shacter, 2000). In particular, the formation of carbonyl groups on proteins can be targeted using the reagent 2,4-dinitrophenylhydrazine (DNPH). This compound reacts with aldehydes to form 2,4-dinitrophenylhydrazone derivatives, which create chromogenic modifications that can be detected at high sensitivity in microplate assays or western blot analysis (Buss et al., 1997; Winterbourn et al., 1999).

In addition, a method involving mass spec analysis to determine carbonyl formation as a result of protein oxidation was developed using a novel mass tag. The carbonyl-specific Element-Coded Affinity Mass Tag (O-ECAT) can covalently couple to aldehyde or ketone oxidation sites using an aminoxy group to form an oxime (Lee et al., 2006; see also Chapter 12, Section 3). The ECAT mass tag consists of a bifunctional metal-chelating group that coordinates a lanthanide metal ion of specific mass. Proteins that have been oxidized to contain carbonyls can be labeled with this reagent and the exact sites of modification determined by analyzing the mass spec signature of the labeled peptides after proteolysis.

Solvent Accessibility of Functional Targets in Proteins

Proteins are highly complex, folded polypeptide chains consisting of at least 20 different amino acids that are strung together in unique sequences, which relate to structure and function. Particular amino acids in proteins may be further modified post-translationally to contain a wide variety of covalent modifications normally found in native proteins. The way in which a peptide chain is wrapped and folded governs each amino acid’s relative exposure to the outside environment, but post-translational modifications can also obscure the protein surface from easy access to the solvent environment.

Amino acid side chains are the primary effectors of the three-dimensional structure of a protein, because their properties vary depending on the presence of charged groups, uncharged polar components, aliphatic chains, aromatic rings, and groups able to form hydrogen bonds with other amino acid residues. The relative hydrophilicity or hydrophobicity of an amino acid side chain is a major factor in determining whether the group will be found on the surface of a globular protein or buried within its globular structure.

However, just considering the individual properties of each amino acid type is not enough to determine its accessibility to the surrounding aqueous environment. There have been many attempts at developing analytical models with predictive value for determining buried or surface accessible amino acids in a folded polypeptide chain (Vasicek et al., 2012). These studies have concluded fractional assignments for each residue that relate to its accessible surface area (ASA) or its solvent exposed area (SEA).

In most cases, there are general trends that emerge from theoretical studies in which hydrophilic amino acids are more likely to be found on the surface of a protein and hydrophobic amino acids are more likely to be inside its three-dimensional structure, but we already knew this intuitively so this conclusion is not surprising. However, a real-life study of the positions of amino acids in proteins whose structures are known is more revealing. The data for Figure 2.21 were calculated from 55 proteins in the Brookhaven database by Bordo and Argos (1991), and the graph was derived from the analysis as presented by the Jena Image Library of Biological Macromolecules (http://www.imb-jena.de/IMAGE_AA.html). Although most of these structures were determined using crystallographic means and thus the proteins are “frozen” in a single structural state, the results are revealing as to how often particular amino acids are accessible to the surrounding solvent.

Figure 2.21. Comparison of the solvent exposed surface area of amino acids in proteins. Data are plotted as a percentage of each amino acid in a protein having greater than a 30-Å2 exposure to the aqueous environment. Charged and polar amino acids are seen to have the most solvent exposure, while uncharged, aromatic, or aliphatic amino acids have the least exposure.

Three levels of SEA are presented in the graph for each amino acid, which corresponds to areas in Å2 accessible to the solvent environment: greater than 30 Å2 for highly accessible amino acids, between 10 Å2 and 30 Å2 for medium accessibility, and less than 10 Å2 for those residues that are relatively not accessible to the solvent. Only the SEA for each amino acid of>30 Å2 is shown in the plotted data. The graph shows that the polar amino acids such as serine, threonine, asparagine, glutamine, and tyrosine often have large areas accessible to the solvent, as do the charged amino acids aspartic acid, glutamic acid, lysine, histidine, and arginine. Surprisingly, proline also falls in the highly accessible group, which is not as expected, because it does not carry a charge, nor is it a highly polar amino acid. However, proline does have a unique characteristic that may explain its appearance on the surface of proteins: it cannot freely rotate about its imino group as other amino acids in a peptide chain can do at amide bonds. This effect results in a kink in the polymer (called a beta-turn), and these sharp turns in a peptide backbone probably occur most often near the surface. Thus, proline is found to be frequently accessible to the solvent environment despite its hydrophobic nature.

The nonpolar amino acids glycine, alanine, valine, leucine, isoleucine, methionine, phenylalanine, and cysteine have lower exposure to the solvent environment than charged or polar residues. However, the frequency at which these groups are found to have an SEA of greater than 30 Å2 is much higher than one would expect based solely upon consideration of their hydrophobicity. In fact, nearly 30 to 50% of the time nonpolar amino acids in a protein can be found at the surface.

At the two extremes, lysine is observed as the amino acid most accessible on the surface of proteins while cysteine is the least exposed amino acid. The inaccessibility of cysteine probably stems from the fact that disulfides are typically buried within the polypeptide structure of proteins, whether they are intrachain or in nature, and proteins rarely contain many reduced cysteine thiols.

It is clear from these data that proteins have complex hydrophilic and hydrophobic regions on their surfaces that determine their potential interactions, binding sites, and active centers. For bioconjugation purposes, targeting of an amino acid even with a high SEA for modification or crosslinking may not result in every residue being modified that is theoretically present in a protein based only on knowledge of its amino acid composition. Even when coupling to very polar or charged groups, such as lysine, there are varying degrees of accessibility to a given reagent, because of the complex folding of the polypeptide chains at the protein surface.

Figure 2.22 shows the globular structure of an immunoglobulin (IgG) Fc region to illustrate this point. In this space-filling model, the lysine residues are highlighted in yellow to easily show their locations within the two polypeptides of the heavy chains. Notice that some of the ε-amino groups at the ends of the side chains are protruding far out into the solvent and are therefore highly accessible for modification. Some of these groups, however, are less exposed even though they are still near the surface, and a few lysines are seen to be between the heavy chain regions where it would be difficult to modify them due to crowding.

Figure 2.22. The solvent accessibility of lysine residues in the Fc region of an antibody is illustrated by highlighting the lysine groups in yellow. Some lysine ε-amine groups are extremely accessible to conjugation, while others are only partially exposed, thus making it difficult to modify all of them in bioconjugation reactions.

Figure 2.64 in this Chapter provides data to validate this effect. The reaction of the thiolating reagent SATA with IgG resulted in only a percentage of the available lysines being modified. As the molar ratio of SATA to IgG was increased, the yield of lysine modification actually became lower. This result can be explained by the relative accessibility of each lysine in the immunoglobulin structure. Some residues are easily accessible and they get modified with high yield even with low molar ratios of SATA-to-IgG. As the molar ratio is increased, it gets more difficult to modify those lysines that are less accessible to the solvent environment or are partially obscured by another polypeptide chain. Thus, the solvent accessibility of particular amino acids is a major factor in whether they can be effectively targeted and modified with a given bioconjugate reagent.

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Peptides are short chains of amino acids connected sequentially by peptide bonds. It is accepted that chains with 2–50 units are classified as peptides, while longer chains are classified as proteins, although the boundary between these two classes of polymers is blurry. Because peptides can be synthesized chemically in large scale as pure standard products or can be fused to carrier proteins and expressed at high levels, they are considered potentially valuable diagnostic probes (Turnbough, 2003). Usually, primarily selected peptides demonstrate modest affinity to target receptors, but their performance may be enhanced by maturation using iterative mutagenesis and selection (Li et al., 1995). It has also been shown that high affinity ligands can be more likely selected from disulfide constrained peptide libraries rather than from linear peptide libraries (O'Neil et al., 1992; reviewed in Smith and Petrenko, 1997). Diagnostic peptide probe technology is still in the infant stages of development, and its prospects may be evaluated after accumulation of a sufficient pool of data. Nevertheless, the promise of this technology may be confirmed by several examples of its successful use, listed in Table 1.

Table 1. Diagnostic and therapeutic probes against biological threats developed using phage display technology

Probe nature//library	Target agent	Analytical test	Sensitivity	Reference
Peptides
	Toxins
Phage//pIII-fused 12-mer peptide library	Staphylococcal enterotoxin B (SEB),	fluoroimmunoassay on the plates	1.4 ng/well	(Goldman et al., 2000)
Peptide//pIII-fused 12-mer peptide library	Botulinum neurotoxins	n.d.	n.d.	(Zdanovsky et al., 2001)
	Viruses
Phage and peptide fused with thioredoxin //pVIII-fused 9-mer peptide library	cucumber mosaic virus (CaMV)	dot blot analysis	5 ng of CaMV	(Gough et al., 1999)
Peptides//7- and 12-mer pIII-fused peptide libraries	spores B. subtilis	FACS with PE-coupled peptides	107 spores	(Knurr et al., 2003)
	Spores
Phage//landscape pVIII-fused 8-mer library	Phytophthora capsici zoospores	coprecipitation	n.d.	(Bishop-Hurley et al., 2002)
Peptides//7- and 12-mer pIII-fused peptide libraries	spores B. anthracis	FACS with PE-coupled peptides	107 spores	(Williams et al., 2003)
Peptides, phage//7- and 12-mer pIII-fused peptide libraries	spores B. anthracis, B. subtilis, Bacillus cereus, Bacillus globigii	FACS with PE-coupled peptides or Alexa-labeled phage; fluorescence microscopy	107 spores	(Turnbough, 2003)
	Bacteria
Peptide, phage//12-mer pIII-fused peptide library	Mycobacterium avium subsp. paratuberculosis	paramagnetic beads capture PCR	10–100 cell/ml	(Stratmann et al., 2002)
Landscape phage//8-mer pVIII-fused peptide library	S. typhimurium	ELISA, Flow cytometry, optical and fluorescence microscopy, transmission and scanning electron microscopy, microbalance	n.d.	This work

Antibodies
	Toxins
scAb, expressed in E. coli//immune library	botulinum toxins	surface plasmon resonance, flow cytometry, enzyme linked immunosorbent assay (ELISA, and hand-held immunochromatographic assay	Kd=2nM	(Emanuel et al., 1996, 2000; Zwick et al., 2001)
Purified scFv //nonimmune libraries	botulinum toxins	SPR	Kd=26–72 nM	(Sheets et al., 1998)
	Viruses
Fab//human immune Fab-library	Human immunodeficiency virus (HIV) type 1	surface plasmon resonance, virus neutralization test	Kd=7.7×10−10M; IC50=10−9−10−10 M	(Barbas et al., 1994; Burton et al., 1991; Zwick et al., 2001)
Fab//human immune Fab-library	Hepatitis B	n.d.	n.d.	(Zebedee et al., 1992)
Fab//simian immune Fab-library	Simian immunodeficiency virus (SIV)	n.d.	n.d.	(Glamann et al., 1998)
Fab//human Fab-library	respiratory syncytial virus	n.d.	n.d.	(Barbas et al., 1992; Crowe et al., 1998)
Fab//human immune Fab-library	Puumala virus	immunoblot	n.d.	(Salonen et al., 1998)
Fab//human immune Fab-library	measles virus	radioimmunoprecipitation assays	n.d.	(de Carvalho Nicacio et al., 2002)
scFv//human immune scFv-library	hepatitis C virus (HCV)	ELISA	n.d.	(Chan et al., 1996; Plaisant et al., 1997)
Fab//human immune Fab-library	Herpes simplex virus	n.d.	n.d.	(De Logu et al., 1998; Sanna et al., 1995)
scFv//human nonimmune scFv-library, affinity maturation	Human cytomegalovirus (HCMV)	competition assay with surface plasmon resonance sensor	Ka=4.3×107 M−1	(Pini et al., 1997)
Fab//human immune Fab-library	Human cytomegalovirus HCMV	n.d.	n.d.	(Williamson et al., 1997)
scFv//human mAb-derived scFv-library	Rabies virus	n.d.	n.d.	(Muller et al., 1997)
Fab//human immune Fab-library	Vaccinia virus	ELISA	n.d.	(Schmaljohn et al., 1999)
Fab//human immune Fab-library	Epstein–Barr virus	n.d.	n.d.	(Bugli et al., 2001)
Fab// human immune Fab-library	Ebola virus	n.d.	n.d.	(Maruyama et al., 1999a,b; Meissner et al., 2002)
	Spores
Phage-fused scFv//Naı̈ve human scFv	spores of the genus Bacillus	fluorescence microscopy, ELISA, competition ELISA	107 cfu/ml (in inhibition test)	(Zhou et al., 2002)
	Bacteria
scFv//mutagenized murine mAb-derived scFv-library	Salmonella serogroup B O-polysaccharide	EIA, SPR	Ka=4×107 M−1	(Deng et al., 1994, 1995)
scFv//mice immune scFv-library	Brucella melitensis	ELISA	n.d.	(Hayhurst et al., 2003)
scFv//semisynthetic scFv-library	M. catarrhalis	immunoblot, ELISA	n.d.	(Boel et al., 1998)
scFv//semisynthetic scFv library	S. suis	n.d.	n.d.	(de Greeff et al., 2000)
scFv//”Griffin 1” human synthetic library	L. monocytogenes	bioelectrochemical sensor	500 cell/ml	(Benhar et al., 2001)
scFv//large human nonimmune scFv-library	C. trachomatis	ELISA, dot-blot, immunoblot and immunocytochemistry	n.d.	(Lindquist et al., 2002)

n.d.—Not defined; all antibody display libraries in the table are pIII-fused.

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Chapters and Articles

Peptide Bonds Link Amino Acids Together in Proteins

Abstract

5.2.2 Peptide bond

Peptides

Peptide bond

Nomenclature

Acid-base properties

Biologically important peptides

Peptide bond

Peptides

4.3 Primary structure

4.3.1 Peptide bond

4.3.2 Determination of primary structure

1.1 Protein Structure and Reactivity

Amino Acids

Nucleophilic Reactions and the pI of Amino Acid Side Chains

Secondary, Tertiary, and Quaternary Structure

Prosthetic Groups, Cofactors, and Post-Translational Modifications

Protecting the Native Conformation and Activity of Proteins

Oxidation of Amino Acids in Proteins and Peptides

Solvent Accessibility of Functional Targets in Proteins

Peptide Therapy Guide Editorial Team

Peptide bond

Biochemistry, Peptide - StatPearls - NCBI Bookshelf - NIH

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Peptide Bond Formation and Hydrolysis