Practical Protein Chemistry - A. Darbre 1989

Determination of the composition of protein oligomers. Preparation of monomers and polypeptide chains
Identification of monomers
Methods for determining the molecular weight of monomers

To determine the Molecular Weight of monomers, sodium dodecyl sulfate-Polyacrylamide gel Electrophoresis (SDS-PAGE) and Gel filtration in both classical and modern (HPLC) variants are employed. While either SDS-PAGE or HPLC is generally preferred, HPLC currently lags behind electrophoresis in its resolution of Proteins with very similar molecular weights (Chapter 6). For proteins containing more than 10% CARBOHYDRATES, Chromatography on agarose gels is recommended, as electrophoresis may yield inaccurate results due to differences in how SDS binds to the polypeptide chain versus the polysaccharide fragments. It is also worth noting that a monomer may consist of several polypeptide chains linked by Disulfide Bonds; therefore, molecular weight determination should be performed both before and after the reduction of S—S bridges.

Molecular weight can also be estimated from quantitative analyses of N-terminal Amino Acids (Section 1.3.1.4). In terms of material consumption (sample amount), this method is equivalent to gel filtration, whereas instrumental analysis requires substantially less material (Chapters 12, 15, and 17).

Statistical analysis of data for 500 proteins has shown that the most probable molecular weight range for monomers is between 10,000 and 60,000 [63]. Indeed, the subunit molecular weights of more than ²/₃ of the proteins studied to date fall within this range. Roughly 50% of these proteins are dimers, 30% are tetramers, and 8% are hexamers.

1.3.1.1. SDS-PAGE. In the presence of SDS, most proteins dissociate into subunits. Subunits composed of two or more polypeptide chains joined by disulfide bonds can undergo further dissociation when treated with reducing agents, such as 2-mercaptoethanol (Section 1.5.1). In general, polypeptide chains bind SDS uniformly—at roughly 1.4 g/g of protein [143]—giving the resulting complex a strong negative charge. The native protein's intrinsic net charge is thus masked, and its charge-to-mass ratio becomes practically constant. Furthermore, SDS induces Conformational Changes in the polypeptide chain, even if its exact absolute configuration remains unknown (whether rod-like, bead-like, an elongated ellipsoid, or a random coil). It has been demonstrated that during PAGE, these complexes migrate in accordance with their molecular weight [182]. The length of the polypeptide chain is determined by comparing the electrophoretic mobility of the complex with that of marker proteins of known molecular weights. In practice, this method is implemented in three variants:

1. A continuous buffer system (typically a phosphate buffer [182]) in a homogeneous gel.

2. A discontinuous buffer system in a homogeneous gel [95].

3. A continuous or discontinuous buffer system in a gradient gel.

Each variant has its own Advantages and disadvantages. General Principles and experimental Procedures for SDS-PAGE are thoroughly discussed in A number of comprehensive publications [3, 35, 73, 183]. In a discontinuous system, samples are pre-concentrated in a stacking gel before entering the resolving gel as narrow bands. In a continuous system, pre-concentration does not occur, and applying a large sample volume results in broad bands by the end of the run. If the sample is applied in a small volume, the results from both systems should be identical. Discrepancies occasionally observed between the two may stem from partial dissociation of the complex during electrophoresis in the discontinuous system.

For various reasons, many proteins bind SDS in abnormally low ratios [164]. This may be due to unusual Amino Acid Composition, the presence of carbohydrates, or because the proteins belong to a group with high positive or negative charges. The electrophoretic mobility of such complexes can also vary depending on single Amino Acid Substitutions or conformational changes within the proteins [40, 46, 126]. Such anomalous behavior can be detected and minimized by performing electrophoresis across gels of varying porosity and constructing a Ferguson plot from the experimental data (see below). In other words, to obtain a reliable estimate of the apparent molecular weight in either a discontinuous or continuous system, measurements must be taken in at least four different gels.

In gradient gel electrophoresis, Polypeptides migrate until they reach a zone with a porosity that prevents their further movement. If all SDS-polypeptide complexes share the same shape, the polyacrylamide concentration in this zone can be taken as a characteristic of that specific molecular weight [97, 99, 101, 137]. Although a complex may partially dissociate and lose some SDS, particularly near equilibrium, this does not affect the final outcome. While sample volume does not influence the width of the protein bands, applying overly large volumes is still discouraged. Reproducibly forming gradients in gradient PAGE can occasionally be challenging, but high-quality precast gels are readily available for this purpose.

Apparatus and Reagents. Polyacrylamide gels are prepared either as cylindrical rods in Glass tubes or as slab gels; the latter format is much more convenient for molecular weight determinations. When using tube electrophoresis, variations in electrophoretic mobility can arise from inconsistencies in the degree of gel polymerization, as well as differences in gel Column length and running conditions. The technique for extracting gels from tubes has been refined in detail [35]; notably, extracting gels with monomer concentrations exceeding 10% can prove difficult. Recommendations regarding equipment and safety precautions when handling reagents are discussed in Section 1.2.1.1.

A 30% stock solution containing 29.2% (w/v) acrylamide and 0.8% (w/v) N,N'-methylenebisacrylamide is filtered and stored at 4 °C (under these conditions, the solution remains stable for several months). Before use, the solution is warmed to 20 °C and degassed under a Water aspirator vacuum.

N,N,N',N'-tetramethylethylenediamine (TEMED) is vacuum-distilled and stored in a dark vessel at 4 °C. High-quality reagents are supplied by Biorad and other vendors. It is recommended to use analytical-grade SDS from BDH, as impurities in the SDS—such as sodium tetradecyl sulfate—can negatively affect band resolution and the overall final result [21, 50, 118]. All Other reagents should be of analytical reagent grade (AR). For extra purification, SDS can be recrystallized from ethanol (600 g dissolved in 3 L of 80% ethanol and treated with activated charcoal).

Polyacrylamide Gels. The properties of the gel depend on the total monomer concentration (acrylamide + N,N'-bisacrylamide) and the proportion of the cross-linking agent (N,N'-methylenebisacrylamide) [34]. Gels are prepared via the free-radical polymerization of monomers in aqueous solution using an ammonium persulfate + TEMED catalyst system.

Once the persulfate is added, the solution is stirred until the crystals dissolve and is quickly transferred into the electrophoresis apparatus. Precautions should be taken to prevent air bubbles from forming in the gel. To achieve this, a short piece of thin tubing (silicone or polyethylene) is attached to the syringe needle, allowing the monomer working solution to be gently layered against the wall of the casting unit. The comb for forming wells is carefully inserted into the solution, any air bubbles trapped on the Teeth are removed, and the solution is left undisturbed until polymerization is complete. Polymerization rates can vary depending on the concentrations of ammonium persulfate and TEMED. On average, polymerization should finish within 10–25 minutes after adding the persulfate. Accelerated or delayed polymerization lacks good reproducibility and may result in a gel with uneven pores.

Class="center">Component ratios for preparing 10 mL of acrylamide gel at various concentrations

Sample preparation. Lyophilized protein samples are dissolved in starting buffer (see below); alternatively, protein solutions are dialyzed against starting buffer for 3–12 hours. Because reaching equilibrium SDS concentration in the sample takes time, dry SDS is added to the protein solution prior to dialysis [to a final concentration of 1% (w/v)]. To ensure complete Denaturation and binding with SDS, samples are incubated at 95–100 °C for 2 minutes. The protein concentration in the sample should be 0.5–1 µg/µL; depending on the gel thickness, 2–10 µL of this solution is loaded into the wells. Sample load can be significantly reduced if detection is to be performed using silver staining. When using a discontinuous buffer system, larger volumes can be loaded if necessary, whereas for a continuous system, dilute samples must be concentrated first. To this end, the protein is precipitated with ammonium sulfate or trichloroacetic acid, and the resulting pellet is redissolved in starting buffer to achieve the optimal concentration (0.5–1 µg/µL).

If a protein is poorly soluble (such as Lipoproteins and Membrane Proteins), 8 M urea or a non-ionic detergent like 1% Triton X-100 is added to the sample, or the SDS concentration is increased to up to 10%. In the latter case, the sample and Buffer solutions must be free of potassium ions or guanidine salts, as these anions react with SDS to form poorly soluble precipitates. Polyacrylamide gels are calibrated using standard marker proteins, kits for which are commercially available from several manufacturers (Pharmacia, Biorad, BDH, Sigma, Calbiochem).

Continuous buffer system with constant polyacrylamide concentration [182]. The Main Components of this system are 0.05 M phosphate buffer and 0.1% SDS.

The stock buffer has the following composition: 0.25 M phosphate (pH 7) + 0.5% SDS + 0.02% sodium azide (13 g NaH₂PO₄·2H₂O + 26 g Na₂HPO₄ + 0.2 g NaN₃ + 5.0 g SDS). Gentle heating is required to dissolve the phosphates; the solution is stored at a Temperature above room temperature.

The electrode buffer is prepared by diluting the stock solution with distilled water at a 1:4 ratio.

Samples are dissolved in a buffer of the following composition: 20 mL stock buffer + 1 g SDS + 10 g glycerol + 0.05 g bromophenol blue + distilled water (up to a total volume of 100 mL). For electrophoresis intended to determine the molecular weight of polypeptide chains with reduced disulfide bonds, the buffer must contain 1% (v/v) 2-mercaptoethanol.

After a brief heating step to boiling, samples are loaded into the gel wells using a micropipette or microsyringe. Electrophoresis is run at 200 V until the dye front approaches the bottom edge of the gel. The duration of the run depends on the gel length and slab geometry. Upon completion, staining, destaining, and photography procedures are carried out (Section 1.2.1.1).

Discontinuous system with constant polyacrylamide concentration [95]. The main components of this system are 0.375 M Tris and 0.1% SDS.

The acrylamide concentration in the resolving gel is 5%–25%, and in the stacking gel (20 mm long) it is 4.5%.

The resolving buffer has the following composition: 1.5 M Tris + 0.4% (w/v) SDS, pH 8.8 (adjusted to the specified pH with HCl).

The stacking gel buffer has the following composition: 0.5 M Tris + 0.4% (w/v) SDS, pH 6.8 (adjusted with HCl).

The sample is dissolved in a buffer of the following composition: 0.0625 M Tris + 2.3% (w/v) SDS + 10% (w/v) glycerol + 0.05% (w/v) bromophenol blue.

When determining the molecular weight of polypeptide chains with reduced disulfide bonds, the buffer contains 1% (v/v) 2-mercaptoethanol or 100 mmol/L dithiothreitol.

The electrode buffer has the following composition: 0.025 M Tris + 0.192 M Glycine + 0.1% (w/v) SDS.

Component ratios for preparing 10 ml of acrylamide resolving gel at various concentrations

The solution is deaerated twice: before adding SDS and (secondly) before adding persulfate and TEMED. After pouring into the system, water or water-saturated isobutanol is carefully layered onto The surface of the solution. Upon completion of polymerization, the upper layer is removed by aspiration, the gel surface is washed with stacking gel buffer, and the stacking gel solution is layered on top. To prepare 10 ml of this solution (4.5% concentration), mix 1.5 ml of stock buffer, 2.5 ml of stacking gel buffer, 6.0 ml of water, 10 mg of ammonium persulfate, and 10 µl of TEMED.

After brief heating to boiling, the samples are loaded into the wells using a microsolsyringe or micropipette. Electrophoresis is carried out at 100 V until the tracking dye leaves the stacking gel, and then at 200 V until the dye front reaches the bottom edge of the gel. The duration of electrophoresis depends on the gel length and chamber geometry, typically lasting about 3 hours.

Continuous or stepped systems in gradient polyacrylamide gels. This technique employs the basic buffer systems discussed previously. The only difference from the preceding systems is that an acrylamide concentration gradient is formed within the gel slab. The gradient is generated using a two-vessel gradient maker and a peristaltic pump [113] or another type of mixing device.

Gradipore and Pharmacia manufacture high-quality precast gels (Section 1.2.1.1). Precast gels do not contain SDS; the detergent is introduced into the gel during a pre-electrophoresis step at 50 mA for 3 hours. To prepare the electrode buffer, the stock solution (containing phosphate and SDS) is diluted with distilled water at a 1:9 ratio. After sample loading, electrophoresis is performed at 50 mA for 3.5 hours. During this time, the protein zones reach an equilibrium position constrained by the pore size.

1.3.1.2. Analysis of Results. As noted in Section 1.2.1, the molecular weights of polypeptide chains are determined using a calibration curve. However, electrophoresis in a gel with a constant concentration (porosity) fails to eliminate errors caused by differences in the free mobility Y₀ of the studied protein and the marker proteins [148, 149]. Therefore, it is recommended to perform electrophoresis in several gels of varying concentrations. In this case, the accuracy of the molecular weight determination increases as the systematic deviation decreases with Changes in the total acrylamide concentration. The obtained results can be analyzed using the empirical Ferguson equation [57]

lgR_t = -K_RT + lgY₀

The retardation coefficient K_R for the test protein and marker proteins is determined from the slope of the plot of lgR_f versus total acrylamide concentration T, while the molecular weight of the test protein is found from the K_R versus molecular weight calibration curve of the marker proteins. For a reliable Determination of k_R, electrophoresis should be carried out in at least four gels of different concentrations (7, 8, 9, and 10%), though seven gels are preferable. Experimental data are processed using statistical analysis Methods [148].

1.3.1.3. Gel Filtration. The General principles of gel filtration of native proteins were discussed in Section 1.2.2; chromatography under denaturing conditions has no major peculiarities. In the presence of Denaturing Agents (guanidine hydrochloride, urea, SDS), the conformation of protein molecules changes, often leading to an increase in hydrodynamic volume. Since Separation is based on molecular size, both the molecular weight range and the calibration curve for a specific gel are altered. For example, under non-denaturing conditions on Sepharose CL-6B, proteins with M of 10⁴–4×10⁶ can be separated; under denaturing conditions in the presence of guanidine hydrochloride, the molecular weight range is 3,000–80,000 [8].

It has been shown that at high concentrations of guanidine-HCl, reduced proteins adopt a random coil conformation. As an empirical METHOD FOR DETERMINING the molecular weights of denatured proteins, chromatography in 6 M guanidine-HCl on agarose gels is recommended,

which are not cross-linked (Sepharose, Ultrogel, Bio-Gel A) [60, 111]. The main drawbacks of this method are its duration (3–5 days) and the poor stability of non-cross-linked agarose in the presence of 6 M guanidine-HCl [110]. Cross-linked agarose gels (such as Sepharose CL-6B) are more stable; in the presence of guanidine-HCl, the properties of such gels remain unchanged for at least a year, and the elution rate can be increased 3- to 4-fold [8, 25, 110]. Gel filtration in the presence of guanidine-HCl is particularly important when analyzing Glycoproteins, because if the protein has a high carbohydrate content, SDS-PAGE yields unreliable results. Chromatography on Sephacryl S-200 is an effective method for determining the molecular weights of small polypeptides with M > 30,000 [18]. High standards of purity are required for guanidine-HCl; analytical-grade reagent from Schwarz-Mann is recommended. If necessary, guanidine-HCl is purified by recrystallization or prepared from guanidine carbonate [127].

When working with most gels, 8 M urea or 0.1% SDS is also used as a component of the elution buffer. However, it has been shown that non-cross-linked agarose gels slowly degrade at high urea concentrations. The elution buffer (0.05 M phosphate or 0.1 M ammonium bicarbonate) typically has a pH of 7–8. When using urea, 0.1 M sodium chloride is added to the buffer to reduce Electrostatic Interactions between the gel matrix and the charged groups of the polypeptide chain. It has been found [47] that cyanate forms spontaneously in aqueous urea solutions:

Cyanate accumulation proceeds at a maximum rate at pH >6, although noticeable amounts of cyanate are also formed in acidic media, even at 0°C [71]. Kinetic studies of the reaction between cyanate and protein amino groups show that cyanate and the amino group react in their unprotonated form According to the equation

This is consistent with the differences in reaction rates between the α-amino group (pK_a ≈ 8) and the ε-amino group (pK_a ≈ 10.7): at pH 7, the reaction rate of the α-amino group is 100 times higher [166]. In the pH ranges below 4 and above 10, carbamylation of amino groups does not occur [174]. In addition to amino groups, cyanate reacts with thiol, carbonyl, imidazole, phosphate, and reactive hydroxyl groups of proteins [166].

It is worth recalling here that intact disulfide bonds can continue to hold a polypeptide chain in a folded state even during denaturation in 6 M guanidine-HCl. As a result, the hydrodynamic volume of the polypeptide chain is reduced compared to the fully denatured state (with cleaved S–S bonds). This effect becomes more pronounced as the polypeptide chain lengthens, the number of disulfide groups increases, and the loop sizes grow. In such cases, a certain caution should be exercised when evaluating molecular weight determination results. For control purposes, chromatographic analysis is performed after reduction and alkylation of the disulfide bridges. The sum of the molecular weights of the individual polypeptide chains should correspond to the total molecular weight of the studied protein. If the alkylation step is omitted after reduction of the S–S bonds, chromatography must be carried out at pH 3 (under conditions where SH-containing chains are sufficiently stable at room temperature) or in the presence of a reducing agent, such as

2-mercaptoethanol (0.1 mol/L). The presence of a reducing agent often interferes with protein detection in the eluate at 280 nm, because the oxidized form of 2-mercaptoethanol absorbs in this region. In such cases, other Analytical Methods are used; for instance, a radioactive label can be introduced into the protein prior to chromatography (Section 1.4.5.1).

Preparation of guanidine hydrochloride [127]: 1 kg of guanidine carbonate is dissolved in 2 L of water at 40 °C, 3 L of ethanol is added, and the mixture is kept in a cold place for several hours. The crystalline mass is collected by filtration, washed twice with aqueous ethanol (ethanol:water = 2:1), and then with ethanol. The recrystallized guanidine carbonate (yield 880 g) is dissolved in 500 mL of water, the solution is cooled in an ice bath, and 6 M HCl is slowly added with stirring until the pH reaches 4 (this pH should be maintained for 12 h). The resulting guanidine hydrochloride is purified by fractional crystallization. A portion of the solution is evaporated at 40 °C until crystallization begins, kept in the cold, and the crystals are separated by filtration; the mother liquor is combined with the next portion of the main solution, and the Procedure is repeated. The final product—780 g of guanidine hydrochloride—can be recrystallized a second time from water according to the described procedure or from methanol. If necessary, activated charcoal is used to remove colored impurities. The optical density of a 6 M guanidine hydrochloride solution at 225 nm should be 0.1.

Preparation of urea solutions. As noted above, the storage of urea solutions leads to The formation of cyanate, with The rate of cyanate formation depending on the solution pH [71]. Therefore, immediately before use, urea solutions must be passed through a mixed-bed ion-exchange column (e.g., an elgalite column). The quality of the urea solution is monitored by measuring electrical conductivity, which should be ≤2 µS. Amines, such as Tris [75] or ethanolamine [41], are sometimes added to urea solutions to bind cyanate.

1.3.1.4. Quantitative analysis of N-terminal amino acids. The determination of N-terminal amino acids is used to demonstrate a high degree of purity of an oligomer (monomer) or individual polypeptide chains. Quantitative analysis of N-terminal amino acids provides insight into the molecular weight of the starting protein or allows the calculation of the number of polypeptide chains. For example, if the content of the N-terminal amino acid is twice the theoretical value or if two different Amino acids are found in an equimolar ratio, it can be concluded that the oligomer consists of two different Peptides. This assumes that the N-terminal amino acids are not blocked, for instance, due to carbamylation in urea solutions. Some amino acids, such as asparagine, glutamine, and Serine, are unstable under Edman Degradation conditions; in such cases, corrections are introduced when interpreting the results.

Typically, the direct Edman method is employed, and PTH-amino acids are quantified by measuring absorbance at 269 nm in ethanol [55]. For the Quantitative determination of PTH-amino acids, prior purification of the modified protein, followed by Hydrolysis and extraction, is recommended.

A procedure has been developed in which the isotope dilution method is combined with Edman degradation [15]. According to one variant of this method, a single step of Edman degradation is performed to block the $\varepsilon$-amino groups of Lysine residues, followed by the Condensation of the second N-terminal residue with a radiolabeled reagent [94]. In this case, there is no need to extract the N-terminal amino acid derivative. Amino acid analysis is used to determine the protein content in the sample, and radioactivity is measured in aliquots of the hydrolyzate. To determine the specific radioactivity of [¹⁴C]FITC, a sample of the purified peptide is modified in a similar manner.

To enhance Protein solubility, sulfophenyl isothiocyanate [23] can be used in the first step, or SDS (1%) can be introduced into the buffer In the second step. If foaming occurs due to the presence of SDS during the benzene extraction step, a drop of n-octanol is added.

Quantitative determination of N-terminal amino acids [15]. The protein content in the sample is determined by amino acid analysis. An accurately weighed sample ($W$ in milligrams) of lyophilized, desalted protein is condensed with FITC of known specific radioactivity, excess reagent is removed, and the Cleavage and cyclization steps are performed. The resulting labeled PTH-amino acid is mixed with a known amount ($N$, in moles) of unlabeled PTH-amino acid, and the specific radioactivity is measured. The molecular weight $M_r$ of the subunit is calculated using the formula

$S_1$ and $S_2$ (Ci$\cdot$mol^-1) are the specific radioactivity of FITC and the PTH-amino acid, respectively.

Edman degradation. To an aliquot (400 µL) of a protein solution (~10 mg/mL) of known concentration, 600 µL of pyridine, 47 µL of N-dimethylallylamine, and ~3 µL of anhydrous trifluoroacetic acid are added, bringing the pH of the reaction mixture to 9.5. Then, 10 µL of phenyl-¹⁵C-isothiocyanate of known specific radioactivity is added, and the mixture is incubated under a nitrogen atmosphere at 40 °C for 1.5 h. Excess reagent is extracted with benzene (2 mL $\times$ 3), and the aqueous phase is lyophilized. To the dry residue, 1 mL of water and glacial acetic acid saturated with dry HCl are added, and the mixture is incubated at 40 °C for 2 h. (For the detailed Edman degradation protocol, see also Chapter 12.)

Isolation and identification of the PTH-amino acid. A known amount of the unlabeled N-terminal amino acid derivative is added to the resulting reaction mixture, which is then extracted with ethyl acetate (2 mL $\times$ 2); the extracts are combined and evaporated to dryness. The PTH-amino acid is isolated by paper chromatography on Whatman SG81 paper using two solvent systems. First, a chloroform–methanol (9:1) system is used; the zone corresponding to THE POSITION OF the PTH-amino acid is eluted with ethyl acetate. The sample is re-chromatographed in chloroform, and the zone is extracted with ethanol. PTH-Amino acids can also be isolated using HPLC (Chapter 13). The concentration of the PTH-amino acid in ethanol is determined by measuring its optical density at 269 nm ($\varepsilon = 16,000\text{ M}^{-1}\cdot\text{cm}^{-1}$) against ethanol. Purity is assessed by the $A_{245}/A_{269}$ ratio (for the reference standard, this ratio is 0.39). Aliquots (~3 µL) are taken to measure radioactivity.

Determination of the specific radioactivity of phenyl isothiocyanate. Labeled FITC (0.5 mCi) is diluted with unlabeled reagent to a specific radioactivity of 0.4 Ci/mol. An aliquot (2 µL) of the reagent is added to a sample of the N-terminal amino acid (2 mg) in 1 mL of aqueous pyridine (1:1) and incubated under a nitrogen atmosphere at 50 °C for 35 min. Excess FITC is extracted with benzene (2 mL $\times$ 4), and the aqueous phase is lyophilized. Cyclization and rearrangement to the phenylthiohydantoin are carried out by incubation in 1 M HCl (500 µL) at 80 °C for 10 min. The PTH-amino acid is extracted with ethyl acetate (2 mL $\times$ 2) and then purified by chromatography on Whatman SG81 paper or by HPLC. The content of the PTH-amino acid is determined from its optical density at 269 nm, after which the number of counts in the aliquot is measured to calculate the specific radioactivity. The average specific radioactivity is calculated based on the analysis of four PTH-amino acid samples.

Scintillation cocktails. To prepare the liquid scintillator, 5 g of 2-(4-tert-butylphenyl)-5-(4-biphenyl)oxadiazole (butyl-PBD) is dissolved in 5 L of toluene (dried over sodium wire). Samples of the PTH-amino acid in 3 mL of ethanol are mixed with 15 mL of liquid scintillator and counted in a radioactivity counter until 40,000 counts are accumulated. The sample radioactivity is calculated using the formula $D_n = N_h/E$, where $N_h$ is the observed count rate and $E$ is the counting efficiency (for the given scintillator and radioactivity counter). $E$ is determined from a calibration curve as $E = N_c/D_c$, where $N_c$ is counts per minute (recorded by the instrument) and $D_c$ is disintegrations per minute of the standard. The counting efficiency for $\beta$-radiation from radioactive carbon is 92–95%.