2. Biophysical and Biochemical Analysis of Recombinant Proteins

Most proteins which are developed for therapy perform specific functions by interacting with other small and large molecules, e.g., cell-surface receptors, binding proteins, nucleic acids, carbohydrates, and lipids. The functional properties of proteins are derived from their folding into distinct three-dimensional structures. Each protein fold is based on its specific polypeptide sequence in which different amino acids are connected through peptide bonds in a specific way. This alignment of the 20 amino acids, called a primary sequence, has in general all the information necessary for folding into a distinct tertiary structure comprising different secondary structures such as α-helices and β-sheets (see below). Because the 20 amino acids possess different side chains, polypeptides with widely diverse properties are obtained.

All of the 20 amino acids consist of a C_α carbon to which an amino group, a carboxyl group, a hydrogen, and a side chain bind in L configuration (Fig. 2.1). These amino acids are joined by condensation to yield a peptide bond consisting of a carboxyl group of an amino acid joined with the amino group of the next amino acid (Fig. 2.2).

The condensation gives an amide group, NH, at the N-terminal side of C_α and a carbonyl group, C = O, at the C-terminal side. These groups, as well as the amino acyl side chains, play important roles in protein folding. Due to their ability to form hydrogen bonds, they make major energetic contributions to the formation of two important secondary structures, α-helix and β-sheet. The peptide bonds between various amino acids are very much equivalent, however, so that they do not determine which part of a sequence should form an α-helix or β-sheet. Sequence-dependent secondary structure formation is determined by the side chains.

The 20 amino acids commonly found in proteins are shown in Fig. 2.3. They are described by their full names and three- and one-letter codes. Their side chains are structurally different in such a way that at neutral pH, aspartic and glutamic acid are negatively charged and lysine and arginine are positively charged. Histidine is positively charged to an extent that depends on the pH. At pH 7.0, on average, about half of the histidine side chains are positively charged. Tyrosine and cysteine are protonated and uncharged at neutral pH, but become negatively charged above pH 10 and 8, respectively.

Polar amino acids consist of serine, threonine, asparagine, and glutamine, as well as cysteine, while nonpolar amino acids consist of alanine, valine, phenylalanine, proline, methionine, leucine, and isoleucine. Glycine behaves neutrally while cystine, the oxidized form of cysteine, is characterized as hydrophobic. Although tyrosine and tryptophan often enter into polar interactions, they are better characterized as nonpolar, or hydrophobic, as described later.

These 20 amino acids are incorporated into a unique sequence based on the genetic code, as the example in Fig. 2.4 shows. This is an amino acid sequence of granulocyte-colony-stimulating factor (G-CSF), which selectively regulates proliferation and maturation of neutrophils. Although the exact properties of this protein depend on the location of each amino acid and hence the location of each side chain in the three-dimensional structure, the average properties can be estimated simply from the amino acid composition, as shown in Table 2.1, i.e., a list of the total number of each type of amino acid contained in this protein molecule.

Using the pK_a values of these side chains and one amino and carboxyl terminus, one can calculate total charges (positive plus negative charges) and net charges (positive minus negative charges) of a protein as a function of pH, i.e., a titration curve. Since cysteine can be oxidized to form a disulfide bond or can be in a free form, accurate calculation above pH 8 requires knowledge of the status of cysteinyl residues in the protein. The titration curve thus obtained is only an approximation, since some charged residues may be buried and the effective pKa values depend on the local environment of each residue. Nevertheless, the calculated titration curve gives a first approximation of the overall charged state of a protein at a given pH and hence its solution property. Other molecular parameters, such as isoelectric point (pI, where the net charge of a protein becomes zero), molecular weight, extinction coefficient, partial specific volume, and hydrophobicity, can also be estimated from the amino acid composition, as shown in Table 2.1.

The primary structure of a protein, i.e., the sequence of the 20 amino acids, can lead to the three-dimensional structure because the amino acids have diverse physical properties. First, each type of amino acid has the tendency to be more preferentially incorporated into certain secondary structures. The frequencies with which each amino acid is found in α-helix, β-sheet, and β-turn, secondary structures that are discussed later in this chapter, can be calculated as an average over a number of proteins whose three-dimensional structures have been solved. These frequencies are listed in Table 2.2. The β-turn has a distinct configuration consisting of four sequential amino acids and there is a strong preference for specific amino acids in these four positions. For example, asparagine has an overall high frequency of occurrence in a β-turn and is most frequently observed in the first and third position of a β-turn. This characteristic of asparagine is consistent with its side chain being a potential site of N-linked glycosylation. Effects of glycosylation on the biological and physicochemical properties of proteins are extremely important. However, their contribution to structure is not readily predictable based on the amino acid composition.

Based on these frequencies, one can predict for particular polypeptide segments which type of secondary structure they are likely to form. As shown in Fig. 2.5a, there are a number of methods developed to predict the secondary structure from the primary sequence of the proteins. Using G-CSF (Fig. 2.5b) as an example, regions of α-helix, β-sheets, turns, hydrophilicity, and antigen sites can be suggested.

Another property of amino acids, which impacts on protein folding, is the hydrophobicity of their side chains. Although nonpolar amino acids are basically hydrophobic, it is important to know how hydrophobic they are. This property has been determined by measuring the partition coefficient or solubility of amino acids in water and organic solvents and normalizing such parameters relative to glycine. Relative to the side chain of glycine, a single hydrogen, such normalization shows how strongly the side chains of nonpolar amino acids prefer the organic phase to the aqueous phase. A representation of such measurements is shown in Table 2.3. The values indicate that the free energy increases as the side chain of tryptophan and tyrosine are transferred from an organic solvent to water and that such transfer is thermodynamically unfavorable. Although it is unclear how comparable the hydrophobic property is between an organic solvent and the interior of protein molecules, the hydrophobic side chains favor clustering together, resulting in a core structure with properties similar to an organic solvent. These hydrophobic characteristics of nonpolar amino acids and hydrophilic characteristics of polar amino acids generate a partition of amino acyl residues into a hydrophobic core and hydrophilic surface, resulting in overall folding.

Combination of the various secondary structures in a protein results in its three-dimensional structure. Many proteins fold into a fairly compact, globular structure.

The folding of a protein molecule into a distinct three-dimensional structure determines its function. Enzyme activity requires the exact coordination of catalytically important residues in the three-dimensional space. Binding of antibody to antigen and binding of growth factors and cytokines to their receptors all require a distinct, specific surface for high-affinity binding. These interactions do not occur if the tertiary structures of antibodies, growth factors, and cytokines are altered.

A unique tertiary structure of a protein can often result in the assembly of the protein into a distinct quaternary structure consisting of a fixed stoichiometry of protein chains within the complex. Assembly can occur between the same proteins or between different polypeptide chains. Each molecule in the complex is called a subunit. Actin and tubulin self-associate into F-actin and microtubule, while hemoglobin is a tetramer consisting of two α- and two β-subunits. Among the cytokines and growth factors, interferon-γ is a homodimer, while platelet-derived growth factor is a homodimer of either A or B chains or a heterodimer of the A and B chain. The formation of a quaternary structure occurs via non-covalent interactions or through disulfide bonds between the subunits.

Interactions occurring between chemical groups in proteins are responsible for formation of their specific secondary, tertiary, and quaternary structures. Either repulsive or attractive interactions can occur between different groups. Repulsive interactions consist of steric hindrance and electrostatic effects. Like charges repel each other and bulky side chains, although they do not repel each other, cannot occupy the same space. Folding is also against the natural tendency to move toward randomness, i.e., increasing entropy. Folding leads to a fixed position of each atom and hence a decrease in entropy. For folding to occur, this decrease in entropy, as well as the repulsive interactions, must be overcome by attractive interactions, i.e., hydrophobic interactions, hydrogen bonds, electrostatic attraction, and van der Waals interactions. Hydration of proteins, discussed in the next section, also plays an important role in protein folding.

These interactions are all relatively weak and can be easily broken and formed. Hence, each folded protein structure arises from a fine balance between these repulsive and attractive interactions. The stability of the folded structure is a fundamental concern in developing protein therapeutics.

Water molecules are bound to proteins internally and externally. Some water molecules occasionally occupy small internal cavities in the protein structure and are hydrogen bonded to peptide bonds and side chains of the protein and often to a prosthetic group, or cofactor, within the protein. The protein surface is large and consists of a mosaic of polar and nonpolar amino acids, and it binds a large number of water molecules, i.e., it is hydrated, from the surrounding environment. As described in the previous section, water molecules trapped in the interior of protein molecules are bound more tightly to hydrogen-bonding donors and acceptors because of a lower dielectric constant.

Solvent around the protein surface clearly has a general role in hydrating peptide and side chains but might be expected to be rather mobile and nonspecific in its interactions. Well-ordered water molecules can make significant contributions to protein stability. One water molecule can hydrogen bond to two groups distant in the primary structure on a protein molecule, acting as a bridge between these groups. Such a water molecule may be highly restricted in motion and can contribute to the stability, at least locally, of the protein, since such tight binding may exist only when these groups assume the proper configuration to accommodate a water molecule that is present only in the native state of the protein. Such hydration can also decrease the flexibility of the groups involved.

There is also evidence for solvation over hydrophobic groups on the protein surface. So-called hydrophobic hydration occurs because of the unfavorable nature of the interaction between water molecules and hydrophobic surfaces, resulting in the clustering of water molecules. Since this clustering is energetically unfavorable, such hydrophobic hydration does not contribute to the protein stability. However, this hydrophobic hydration facilitates hydrophobic interaction. This unfavorable hydration is diminished as the various hydrophobic groups come in contact either intramolecularly or intermolecularly, leading to the folding of intrachain structures or to protein-protein interactions.

Both the loosely and strongly bound water molecules can have an important impact, not only on protein stability but also on protein function. For example, certain enzymes function in nonaqueous solvent provided that a small amount of water, just enough to cover the protein surface, is present. Bound water can modulate the dynamics of surface groups, and such dynamics may be critical for enzyme function. Dried enzymes are, in general, inactive and become active after they absorb 0.2 g water per g protein. This amount of water is only sufficient to cover surface polar groups, yet may give sufficient flexibility for function.

Evidence that water bound to protein molecules has a different property from bulk water can be demonstrated by the presence of non-freezable water. Thus, when a protein solution is cooled below −40 °C, a fraction of water, ~0.3 g water/g protein, does not freeze and can be detected by high-resolution NMR. Several other techniques also detect a similar amount of bound water. This unfreezable water reflects the unique property of bound water that prevents it from adopting an ice structure.

Proteins are immersed under physiological conditions or in test tubes in aqueous solutions containing not only water but also other solution components, e.g., salts, metals, amino acids, sugars, and many other minor components. These components also interact with the protein surface and affect protein folding and stability. For examples, sugars and amino acids are known to enhance folding and stability of the proteins, as described below.

Conventional spectroscopic techniques used to obtain information on the folded structure of proteins are circular dichroism (CD), fluorescence, and Fourier transform infrared spectroscopies (FTIR). CD and FTIR are widely used to estimate the secondary structure of proteins. The α-helical content of a protein can be readily estimated by CD in the far-UV region (180–260 nm) and by FTIR. FTIR signals from loop structures, however, occasionally overlap with those arising from an α-helix. The β-sheet gives weak CD signals, which are variable in peak positions and intensities due to twists of interacting β-strands, making far-UV CD unreliable for evaluation of these structures. On the other hand, FTIR can reliably estimate the β-structure content as well as distinguish between parallel and antiparallel forms.

CD in the near-UV region (250–340 nm) reflects the environment of aromatic amino acids, i.e., tryptophan, tyrosine, and phenylalanine, as well as that of disulfide structures. Fluorescence spectroscopy yields information on the environment of tyrosine and tryptophan residues. CD and fluorescence signals in many cases are drastically altered upon refolding and hence can be used to follow the formation of the tertiary structure of a protein.

None of these techniques can give the folded structure at the atomic level, i.e., they give no information on the exact location of each amino acyl residue in the three-dimensional structure of the protein. This information can only be determined by X-ray crystallography or NMR. However, CD, FTIR, and fluorescence spectroscopic methods are fast and require lower protein concentrations than either NMR or X-ray crystallography and are amenable for the examination of the protein under widely different conditions. When a naturally occurring form of the protein is available, these techniques, in particular near-UV CD and fluorescence spectroscopies, can quickly address whether the refolded protein assumes the native folded structure.

Temperature dependence of these spectroscopic properties also provides information about protein folding. Since the folded structures of proteins are built upon cooperative interactions of many side chains and peptide bonds in a protein molecule, elimination of one interaction by heat can cause cooperative elimination of other interactions, leading to the unfolding of protein molecules. Thus, many proteins undergo a cooperative thermal transition over a narrow temperature range. Conversely, if the proteins are not fully folded, they may undergo noncooperative thermal transitions as observed by a gradual signal change over a wider range of temperature.

Such a cooperative structure transition can also be examined by differential scanning calorimetry. When the structure unfolds, it requires heat. Such heat absorption can be determined using this highly sensitive calorimetry technique.

Hydrodynamic properties of proteins change greatly upon folding, going from elongated and expanded structures to compact globular ones. Sedimentation velocity and size-exclusion chromatography (see section “Analytical Techniques”) are two frequently used techniques for the evaluation of hydrodynamic properties, although the latter is much more accessible. The sedimentation coefficient (how fast a molecule migrates in a centrifugal field) is a function of both the molecular weight and hydrodynamic size of the proteins, while elution position in size-exclusion chromatography (how fast it migrates through pores) depends only on the hydrodynamic size (see Chap.​ 3). In both methods, comparison of the sedimentation coefficient or elution position with that of a globular protein with an identical molecular weight (or upon appropriate molecular-weight normalization) gives information on how compactly the protein is folded.

For oligomeric proteins, the determination of molecular weight of the associated states and acquisition of the quaternary structure can be used to assess the folded structure. For strong interactions, specific protein association requires that intersubunit contact surfaces perfectly match each other. Such an associated structure, if obtained by covalent bonding, may be determined simply by sodium dodecyl sulfate-polyacrylamide gel electrophoresis. If protein association involves non-covalent interactions, sedimentation equilibrium or light scattering experiments can assess this phenomenon. Although these techniques have been used for many decades with some difficulty, emerging technologies in analytical ultracentrifugation and laser light scattering, and appropriate software for analyzing the results, have greatly facilitated their general use, as described in detail below.

Two fundamentally different light scattering techniques can be used in characterizing recombinant proteins. “Static” light scattering measures the intensity of the scattered light. “Dynamic” light scattering measures the fluctuations in the scattered light intensity as molecules diffuse in and out of a very small scattering region (Brownian motion).

Static light scattering is often used online in conjunction with size-exclusion chromatography (SEC). The scattering signal is proportional to the product of molecular mass times weight concentration. Dividing this signal by one proportional to the concentration, such as obtained from an UV absorbance or refractive index detector, then gives a direct and absolute measure of the mass of each peak eluting from the column, independent of molecular conformation and elution position. This SEC-static scattering combination allows rapid identification of whether the native state of a protein is a monomer or an oligomer and the stoichiometry of multi-protein complexes. It is also very useful in identifying the mass of aggregates which may be present and thus is useful for evaluating protein stability.

Dynamic light scattering (DLS) measures the diffusion rate of the molecules, which can be translated into the Stokes radius, a measure of hydrodynamic size. Although the Stokes radius is strongly correlated with molecular mass, it is also strongly influenced by molecular shape (conformation), and thus, DLS is far less accurate than static scattering for measuring molecular mass. The great strength of DLS is its ability to cover a very wide size range in one measurement and to detect very small amounts of large aggregates (<0.01 % by weight). Other important advantages over static scattering with SEC are a wide choice of buffer conditions and no potential loss of species through sticking to a column.

An analytical ultracentrifuge incorporates an optical system and special rotors and cells in a high-speed centrifuge to permit measurement of the concentration of a sample versus position within a spinning centrifuge cell. There are two primary strategies: analyzing either the sedimentation velocity or the sedimentation equilibrium. When analyzing the sedimentation velocity, the rotor is spun at very high speed, so the protein sample will completely sediment and form a pellet. The rate at which the protein pellets is measured by the optical system to derive the sedimentation coefficient, which depends on both mass and molecular conformation. When more than one species is present (e.g., a monomer plus a covalent dimer degradation product), a separation is achieved based on the relative sedimentation coefficient of each species.

Proteins form not only small oligomers that can be measured by the above techniques but also much larger aggregates, called subvisible and visible particles, which are present in minute quantities. As their size approaches the size of virus, they become highly immunogenic (cf. Chap.​ 6) and hence determination of their size and amount becomes critical for developing pharmaceutical protein products. Such determination requires imaging of the particles, as the hydrodynamic techniques such as dynamic light scattering and sedimentation velocity have neither sensitivity nor resolution for such large aggregates heterogeneous in size distribution. Normally such particles are present in minute quantity, but yet cause serious immunogenic responses due to their large size.

Because the sedimentation coefficient is sensitive to molecular conformation and can be measured with high precision (~0.5 %), sedimentation velocity can detect even fairly subtle differences in conformation. This ability can be used, for example, to confirm that a recombinant protein has the same conformation as the natural wild-type protein or to detect small changes in structure with changes in the pH or salt concentration that may be too subtle to detect by other techniques, such as CD or differential scanning calorimetry.

In sedimentation equilibrium, a much lower rotor speed and milder centrifugal force is used than for sedimentation velocity. The protein still accumulates toward the outside of the rotor, but no pellet is formed. This concentration gradient across the cell is continuously opposed by diffusion, which tries to restore a uniform concentration. After spinning for a long time (usually 12–36 h), an equilibrium is reached where sedimentation and diffusion are balanced and the distribution of protein no longer changes with time. At sedimentation equilibrium, the concentration distribution depends only on the molecular mass and is independent of molecular shape. Thus, self-association for the formation of dimers or higher oligomers (whether reversible or irreversible) is readily detected, as are binding interactions between different proteins. For reversible association, it is possible to determine the strength of the binding interaction by measuring samples over a wide range of protein concentrations.

In biotechnology applications, sedimentation equilibrium is often used as the “gold standard” for confirming that a recombinant protein has the expected molecular mass and biologically active state of oligomerization in solution. It can also be used to determine the average amount of glycosylation or conjugation of moieties such as polyethylene glycol. The measurement of binding affinities for receptor-cytokine, antigen-antibody, or other interaction can also sometimes serve as a functional characterization of recombinant proteins (although some of these interactions are too strong to be measured by this method).

Site-specific chemical modification and proteolytic digestion are also powerful techniques for studying the folding of proteins. The extent of chemical modification or proteolytic digestion depends on whether the specific sites are exposed to the solvent or are buried in the interior of the protein molecules and are thus inaccessible to these modifications. For example, trypsin cleaves peptide bonds on the C-terminal side of basic residues. Although most proteins contain several basic residues, brief exposure of the native protein to trypsin usually generates only a few peptides, as cleavage occurs only at the accessible basic residues, whereas the same treatment can generate many more peptides when done on the denatured (unfolded) protein, since all the basic residues are now accessible (see also peptide mapping in section “Mass Spectrometry”).

Blotting methods form an important niche in biotechnology. They are used to detect very low levels of unique molecules in a milieu of proteins, nucleic acids, and other cellular components. They can detect aggregates or breakdown products occurring during long-term storage and they can be used to detect components from the host cells used in producing recombinant proteins.

Biomolecules are transferred to a membrane (“blotting”), and this membrane is then probed with specific reagents to identify the molecule of interest. Membranes used in protein blots are made of a variety of materials including nitrocellulose, nylon, and polyvinylidene difluoride (PVDF), all of which avidly bind protein.

Liquid samples can be analyzed by methods called dot blots or slot blots. A solution containing the biomolecule of interest is filtered through a membrane which captures the biomolecule. The difference between a dot blot and a slot blot is that the former uses a circular or disk format, while the latter is a rectangular configuration. The latter method allows for a more precise quantification of the desired biomolecule by scanning methods and relating the integrated results to that obtained with known amounts of material.

Often, the sample is subjected to some type of fractionation, such as polyacrylamide gel electrophoresis, prior to the blotting step. An early technique, Southern blotting, named after the discoverer, E.M. Southern, is used to detect DNA fragments. When this procedure was adapted to RNA fragments and to proteins, other compass coordinates were chosen as labels for these procedures, i.e., northern blots for RNA and western blots for proteins. Western blots involve the use of labeled antibodies to detect specific proteins.

Analytical methodologies for measuring protein properties stem from those used in their purification. The major difference is that systems used for analysis have a higher resolving power and lower detection limit than those used in purification. The two major methods for analysis have their bases in chromatographic or electrophoretic techniques.

Chromatography techniques are used extensively in biotechnology not only in protein purification procedures (see Chap.​ 3) but also in assessing the integrity of the product. Routine procedures are highly automated so that comparisons of similar samples can be made. An analytical system consists of an autosampler which will take a known amount (usually a known volume) of material for analysis and automatically places it in the solution stream headed toward a separation column used to fractionate the sample. Another part of this system is a pump module which provides a reproducible flow rate. In addition, the pumping system can provide a gradient which changes properties of the solution such as pH, ionic strength, and hydrophobicity. A detection system (or possibly multiple detectors in series) is located at the outlet of the column. This measures the relative amount of protein exiting the column. Coupled to the detector is a data acquisition system which takes the signal from the detector and integrates it into a value related to the amount of material (see Fig. 2.12). When the protein appears, the signal begins to increase, and as the protein passes through the detector, the signal subsequently decreases. The area under the peak of the signal is proportional to the amount of material which has passed through the detector. By analyzing known amounts of protein, an area versus amount of protein plot can be generated and this may be used to estimate the amount of this protein in the sample under other circumstances. Another benefit of this integrated chromatography system is that low levels of components which appear over time can be estimated relative to the major desired protein being analyzed. This is a particularly useful function when the long-term stability of the product is under evaluation.

Chromatographic systems offer a multitude of different strategies for successfully separating protein mixtures and for quantifying individual protein components (see Chap.​ 3). The following describes some of these strategies.

Paramount to the development of a protein therapeutic is to have an assay that identifies its biological function. Chromatographic and electrophoretic methodologies can address the homogeneity of a biotherapeutic and be useful in investigating stability parameters. However, it is also necessary to ascertain whether the protein has acceptable bioactivity. Bioactivity can be determined either in vivo, i.e., by administering the protein to an animal and ascertaining some change within its body (function), or in vitro. Bioassays in vitro monitor the response of a specific receptor or microbiological or tissue cell line when the therapeutic protein is added to the system. An example of an in vitro bioassay is the increase in DNA synthesis in the presence of the therapeutic protein as measured by the incorporation of radioactively labeled thymidine. The protein factor binds to receptors on the cell surface that triggers secondary messengers to send signals to the cell nucleus to synthesize DNA. The binding of the protein factor to the cell surface is dependent upon the amount of factor present. Figure 2.14 presents a dose–response curve of thymidine incorporation as a function of concentration of the factor. At low concentrations, the factor is too low to trigger a response. As the concentration increases, the incorporation of thymidine occurs, and at higher concentrations, the amount of thymidine incorporation ceases to increase as DNA synthesis is occurring at the maximum rate. A standard curve can be obtained using known quantities of the protein factor. Comparison of other solutions containing unknown amounts of the factor with this standard curve will then yield quantitative estimates of the factor concentration. Through experience during the development of the protein therapeutic, a value is obtained for a fully functional protein. Subsequent comparisons to this value can be used to ascertain any loss in activity during stability studies or changes in activity when amino acyl residues of the protein are modified.

Other in vitro bioassays can measure changes in cell number or production of another protein factor in response to the stimulation of cells by the protein therapeutic. The amount of the secondary protein produced can be estimated by using an ELISA.

Recent advances in the measurement of the molecular masses of proteins have made this technique an important analytical tool. While this method was used in the past to analyze small volatile molecules, the molecular weights of highly charged proteins with masses of over 100 kilodaltons (kDa) can now be accurately determined.

Because of the precision of this method, posttranslational modifications such as acetylation or glycosylation can be predicted. The masses of new protein forms that arise during stability studies provide information on the nature of this form. For example, an increase in mass of 16 Da suggests that an oxygen atom has been added to the protein as happens when a methionyl residue is oxidized to a methionyl sulfoxide residue. The molecular mass of peptides obtained after proteolytic digestion and separation by HPLC indicates from which region of the primary structure they are derived. Such HPLC chromatogram is called a “peptide map.” An example is shown in Fig. 2.15. This is obtained by digesting a protein with pepsin and by subsequently separating the digested peptides by reverse HPLC. This highly characteristic pattern for a protein is called a “protein fingerprint.” Peaks are identified by elution times on HPLC. If peptides have molecular masses differing from those expected from the primary sequence, the nature of the modification to that peptide can be implicated. Moreover, molecular mass estimates can be made for peptides obtained from unfractionated proteolytic digests. Molecular masses that differ from expected values indicate that a part of the protein molecule has been altered, that glycosylation or another modification has been altered, or that the protein under investigation still contains contaminants.

Another way that mass spectrometry can be used as an analytical tool is in the sequencing of peptides. A recurring structure, the peptide bond, in peptides tends to yield fragments of the mature peptide which differ stepwise by an amino acyl residue. The difference in mass between two fragments indicates the amino acid removed from one fragment to generate the other. Except for leucine and isoleucine, each amino acid has a different mass and hence a sequence can be read from the mass spectrograph. Stepwise removal can occur from either the amino terminus or carboxy terminus.

By changing three basic components of the mass spectrometer, the ion source, the analyzer, and the detector, different types of measurement may be undertaken. Typical ion sources which volatilize the proteins are electrospray ionization, fast atom bombardment, and liquid secondary ion. Common analyzers include quadrupole, magnetic sector, and time-of-flight instruments. The function of the analyzer is to separate the ionized biomolecules based on their mass-to-charge ratio. The detector measures a current whenever impinged upon by charged particles. Electrospray ionization (El) and matrix-assisted laser desorption (MALDI) are two sources that can generate high-molecular-weight volatile proteins. In the former method, droplets are generated by spraying or nebulizing the protein solution into the source of the mass spectrometer. As the solvent evaporates, the protein remains behind in the gas phase and passes through the analyzer to the detector. In MALDI, proteins are mixed with a matrix which vaporizes when exposed to laser light, thus carrying the protein into the gas phase. An example of MALDI-mass analysis is shown in Fig. 2.16, indicating the singly charged ion (116,118 Da) and the doubly charged ion (5,8036.2) for a purified protein. Since proteins are multi-charge compounds, a number of components are observed representing mass-to-charge forms, each differing from the next by one charge. By imputing various charges to the mass-to-charge values, a molecular mass of the protein can be estimated. The latter step is empirical since only the mass-to-charge ratio is detected and not the net charge for that particular particle.