Practical Protein Chemistry - A. Darbre 1989

Peptide mapping of proteins
Peptide mapping in practice
Cleveland mapping

The target protein (labeled or unlabeled) can be isolated from a mixture using traditional Biochemical Methods (such as ammonium sulfate precipitation or open-Column chromatography) or more modern approaches (e.g., immunoprecipitation [13] and Affinity Chromatography [18, 29]). However, purifying a protein to homogeneity by these methods is often difficult, making it necessary at The final stage to use the highest-resolution system available: SDS-Polyacrylamide gel Electrophoresis (SDS-PAGE). In the original version of Cleveland's method [12], Proteins in the gel were lightly stained with Coomassie Brilliant Blue [14], followed by rapid destaining and excision of the corresponding wet gel band.

For radioactively labeled proteins, the gel is removed from the electrophoresis apparatus, washed with ~20 volumes of distilled Water to remove buffer components, and then dried without prior fixation (if the gel has been fixed in acetic acid or methanol, eluting the intact protein from the rehydrated gel is often difficult). The dried gel is then subjected to autoradiography, and the desired protein band is excised using the X-ray film as a template.

A second SDS gel is prepared with an acrylamide concentration such that the mobility of the target protein is ~0.2 relative to the dye front (bromophenol blue) (Fig. 7.1). Gels prepared with an acrylamide gradient are also used; these are particularly useful when the size of the resulting fragments is difficult to predict. The stacking gel should have a relatively low acrylamide content—so as not to restrict the mobility of the intact protein or proteolytic enzyme (if used)—and sufficient length to allow the sample to form a compact band before reaching the resolving gel. The wells must be wide and deep enough to accommodate the rehydrated protein gel slices and the enzyme solution, ensuring free contact between them.

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FIG. 7.1. SDS-PAGE mapping of Acetylcholine Receptor subunits. a — Digestion with V8 protease: lanes 1–4, 5 µg of α-, β-, γ-, and δ-subunits, respectively; lane 5, marker proteins; lanes 6–9 (α), 10–13 (β), 14–17 (γ), and 18–21 (δ) subunits digested with 0.0005, 0.005, 0.05, and 0.25 µg of V8 protease, respectively. b — digestion with Papain: lanes 1–4, 5 µg of α-, β-, γ-, and δ-subunits; lane 5, marker proteins; lanes 6–9 (α), 10–13 (β), 14–17 (γ), and 18–21 (δ) subunits digested with 0.0001, 0.001, 0.01, and 0.1 µg of papain, respectively. Arrows indicate bands treated with the Schiff reagent for carbohydrate detection [32]. (Reprinted with permission from the American Chemical Society.)

The choice of Cleavage method depends on The Nature of the substrate protein and is determined by the experimenter based on experience. Proteolytic Enzymes are widely used, notably the V8 protease isolated from culture filtrates of Staphylococcus aureus [15]. This enzyme cleaves peptide chains at the C-terminal side of glutamic and aspartic acid residues; however, in an appropriate buffer, Hydrolysis is restricted to glutamic acid residues [34]. In standard Laemmli discontinuous buffer systems [40], V8 protease cleaves peptide bonds formed by both dicarboxylic Amino Acids. Because hydrolysis takes place within the stacking gel, the enzyme must retain activity in a 0.1% (w/v) SDS solution, and preferably at higher detergent concentrations as well. Under these conditions, V8 protease [4, 30], papain [32], Elastase, α-Chymotrypsin [29], and Trypsin [4] remain active and can be used.

If the substrate protein is in solution, the enzyme is added directly to it, and the mixture is incubated for a specified time until the desired degree of digestion is achieved. The hydrolyzate is then loaded onto the gel and separated in the usual manner [30, 31]. There is no need to inactivate the protease, as it will be separated from the substrate within the resolving gel. Typically, a reducing agent is added after digestion is complete, and the sample is heated at 100 °C for 2–5 min. This Procedure promotes sharper electrophoretic bands while simultaneously inactivating the enzyme.

More frequently, the protein is digested directly within the polyacrylamide gel: the enzyme solution (optimally ~10 µL) is placed into a well that already contains electrode buffer. The COMPOSITION OF THE enzyme solution should match that of the stacking gel, but with a slightly higher density, which is achieved by adding 10–20% glycerol or sucrose. The gel slice containing the substrate protein is carefully immersed into the well above the protease solution so as not to disturb the sharp boundary. Although it is helpful to use thicker spacers for the second gel than for the first, gel slices can usually be placed into the wells of a gel plate of the same size, provided a tight fit is achieved to prevent the gel slice from dropping into the enzyme solution and splashing. Positioning the substrate gel like a "cushion" atop the denser enzyme solution facilitates more efficient digestion by ensuring an even distribution of the protease across the entire track surface.

When voltage is applied, the substrate protein and the protease migrate and concentrate into narrow, adjacent zones; the current is then switched off (usually for 30–60 min) to allow enzymatic digestion to proceed. The active, native protease cleaves some of the most "susceptible" bonds in the denatured, unfolded substrate. Voltage is then reapplied, and the resulting fragments are separated in the resolving gel.

Several methods are used to detect Peptides on SDS-polyacrylamide gels. A radioactive label can be pre-introduced into the protein (via cellular METABOLISM or in vitro), in which case the separated fragments are detected by autoradiography or fluorography [2]. Specific identification is achieved using an affinity label (such as a natural Ligand or its synthetic analogs) [29] or via enzymes, such as Kinases capable of incorporating radioactive groups (provided the fragments act as substrates). More generally, labeling is accomplished by iodinating accessible Tyrosine residues [11, 53, 54, 45] or metabolically in tissue [13] or organ [59] cultures maintained in a medium containing one or more radiolabeled amino acids. Specific labeling introduces radioactivity into only certain peptide fragments; this approach is particularly informative for investigating local Homology, such as the microenvironment of a ligand-binding site [29] or phosphorylation sites.

General staining Reagents are also used to visualize gels, including Coomassie Brilliant Blue [14] and silver staining complexes [47, 48, 52], as well as more specific ones—such as the Schiff reagent [32] or a modified silver staining protocol [16] for identifying glycopeptides [17, 28, 69], or radio- [39] and fluorescently labeled Lectins [23] (see also Chapter 8).

Polypeptides can be transferred from the gel onto nitrocellulose sheets [7, 25, 67], positively charged membranes [24], or diazotized paper [10, 56] and detected by specific binding to labeled Antibodies [29] or lectins [27, 33]. Antiserum allows the identification of all protein fragments, whereas Monoclonal Antibodies or lectins specifically bind to a single fragment or a limited number of fragments. Because antibodies can detect an individual protein within a mixture, this technique enables the comparison of two proteins, even when one of them has not previously been isolated in pure form. Cleveland mapping is typically used for relatively large peptides, making the application of immunological detection methods highly advantageous due to the high probability that they retain uncleaved linear antigenic determinants.

To obtain an optimal set of protein fragments by proteolysis (in terms of component size), various enzyme loads are tested at a fixed Substrate Concentration [12] (Fig. 7.1). In the absence of data regarding the proteolytic susceptibility of the protein under study (which can vary widely), one starts with an enzyme load of 1 µg per gel well. This amount is then decreased or increased tenfold, depending on the available protein supply. The optimal experimental conditions are subsequently selected based on the results obtained.

Chemical cleavage methods of the polypeptide chain can also be employed: Treatment with Cyanogen bromide (at Methionine residues) [9, 43, 50], formic acid (at aspartic acid–Proline bonds) [41, 58], or N-chlorosuccinimide (at Tryptophan residues) [42]. In these cases, the protein-containing gel slice is treated with the reagent in a tube, and the resulting mixture of fragments is then loaded onto a second gel for Separation. The advantage of chemical methods is obvious when working with small amounts of unlabeled protein that require high-sensitivity silver staining for visualization, which is unaffected by the presence of proteases.

Numerous variations of peptide mapping in gels have been published. Primary gel electrophoresis can be used to separate several proteins, after which a gel strip is excised, laid onto a second gel, subjected to enzymatic digestion as described above, and the resulting fragments are separated by a second round of electrophoresis.

Thus, peptide maps of several proteins present in a mixture can be obtained with minimal effort (provided they exhibit different relative mobilities in the primary gel) [4, 41, 65]. Complications arising from this technique are related to the fact that the concentrations of components in the initial mixture may vary, while digestion is carried out at a single Enzyme Concentration. This can lead to unequal degrees of Protein Cleavage and erroneous Conclusions when analyzing the electrophoregram.

If a protein is available only in very small quantities, its fragments can be transferred electrophoretically from the gel onto diazotized paper, to which they bind covalently, or onto nitrocellulose, where non-covalent attachment takes place. In the presence of appropriate antisera or monoclonal antibodies, protein fragments can be detected on the replica with high sensitivity. For instance, a mixture of hydrolyzates of acetylcholine receptor subunits from three different animal species was successfully separated by one-dimensional electrophoresis [29], and the polypeptides were then transferred to paper containing diazophenylthioether groups [56, 63]. Antisera and monoclonal antibodies directed against each individual receptor subunit were used to detect fragments of a specific subunit within the mixture, considerably simplifying their comparative analysis (Fig. 7.2, a–g). Although the peptide maps differed, monoclonal antibody staining on map g revealed a fragment with identical mobility in all three tracks (4, 5, and 6). This indicated that the receptors from each species share a degree of homology, manifested by the generation of a fragment of identical size with the same linear antigenic determinant in all three cases.

There is no doubt that Cleveland mapping will continue to evolve and improve. Even now, for example, experiments are becoming widespread in which cDNA is synthesized from very low-Abundance cellular mRNAs, cloned, and sequenced. Polyclonal and monoclonal antibodies raised against synthetic peptides deduced from The nucleotide sequence are then used to identify the corresponding proteins on electrophoregrams.