Methods used for amino acid analysis are usually based on a chromatographic separation of the amino acids present in the test sample. Current techniques take advantage of the automated chromatographic instrumentation designed for analytical methodologies. An amino acid analysis instrument will typically be a low-pressure or high-pressure liquid chromatograph capable of generating mobile phase gradients that separate the amino acid analytes on a chromatographic column. The instrument must have postcolumn derivatization capability, unless the sample is analyzed using precolumn derivatization. The detector is usually a UV-visible or fluorescence detector depending on the derivatization method used. A recording device (e.g., integrator) is used for transforming the analog signal from the detector and for quantitation. It is preferred that instrumentation be dedicated particularly for amino acid analysis.

Background contamination is always a concern for the analyst in performing amino acid analysis. High-purity reagents are necessary (e.g., low-purity hydrochloric acid can contribute to glycine contamination). Analytical reagents are changed routinely every few weeks using only high-pressure liquid chromatography (HPLC) grade solvents. Potential microbial contamination and foreign material that might be present in the solvents are reduced by filtering solvents before use, keeping solvent reservoirs covered, and not placing amino acid analysis instrumentation in direct sunlight.

Laboratory practices can determine the quality of the amino acid analysis. Place the instrumentation in a low traffic area of the laboratory. Keep the laboratory clean. Clean and calibrate pipets according to a maintenance schedule. Keep pipet tips in a covered box; the analysts may not handle pipet tips with their hands. The analysts may wear powder-free latex or equivalent gloves. Limit the number of times a test sample vial is opened and closed because dust can contribute to elevated levels of glycine, serine, and alanine.

A well-maintained instrument is necessary for acceptable amino acid analysis results. If the instrument is used on a routine basis, it is to be checked daily for leaks, detector and lamp stability, and the ability of the column to maintain resolution of the individual amino acids. Clean or replace all instrument filters and other maintenance items on a routine schedule.

Acceptable amino acid standards are commercially available1 for amino acid analysis and typically consist of an aqueous mixture of amino acids. When determining amino acid composition, protein or peptide standards are analyzed with the test material as a control to demonstrate the integrity of the entire procedure. Highly purified bovine serum albumin has been used as a protein standard for this purpose.

Calibration of amino acid analysis instrumentation typically involves analyzing the amino acid standard, which consists of a mixture of amino acids at a number of concentrations, to determine the response factor and range of analysis for each amino acid. The concentration of each amino acid in the standard is known. In the calibration procedure, the analyst dilutes the amino acid standard to several different analyte levels within the expected linear range of the amino acid analysis technique. Then, replicates at each of the different analyte levels can be analyzed. Peak areas obtained for each amino acid are plotted versus the known concentration for each of the amino acids in the standard dilution. These results will allow the analyst to determine the range of amino acid concentrations where the peak area of a given amino acid is an approximately linear function of the amino acid concentration. It is important that the analyst prepare the samples for amino acid analysis so that they are within the analytical limits (e.g., linear working range) of the technique employed in order to obtain accurate and repeatable results.

Four to six amino acid standard levels are analyzed to determine a response factor for each amino acid. The response factor is calculated as the average peak area or peak height per nmol of amino acid present in the standard. A calibration file consisting of the response factor for each amino acid is prepared and is used to calculate the concentration of each amino acid present in the test sample. This calculation involves dividing the peak area corresponding to a given amino acid by the response factor for that amino acid to give the nmol of the amino acid. For routine analysis, a single-point calibration may be sufficient; however, the calibration file is updated frequently and tested by the analysis of analytical controls to ensure its integrity.

Consistent high quality amino acid analysis results from an analytical laboratory require attention to the repeatability of the assay. During analysis of the chromatographic separation of the amino acids or their derivatives, numerous peaks can be observed on the chromatogram that corresponds to the amino acids. The large number of peaks makes it necessary to have an amino acid analysis system that can repeatedly identify the peaks based on retention time and integrate the peak areas for quantitation. A typical repeatability evaluation involves preparing a standard amino acid solution and analyzing many replicates (i.e., six analyses or more) of the same standard solution. The relative standard deviation (RSD) is determined for the retention time and integrated peak area of each amino acid. An evaluation of the repeatability is expanded to include multiple assays conducted over several days by different analysts. Multiple assays include the preparation of standard dilutions from starting materials to determine the variation due to sample handling. Often, the amino acid composition of a standard protein (e.g., bovine serum albumin) is analyzed as part of the repeatability evaluation. By evaluating the replicate variation (i.e., RSD), the laboratory can establish analytical limits to ensure that the analyses from the laboratory are under control. It is desirable to establish the lowest practical variation limits to ensure the best results. Areas to focus on to lower the variability of the amino acid analysis include sample preparation, high background spectral interference due to quality of reagents and/or laboratory practices, instrument performance and maintenance, data analysis and interpretation, and analyst performance and habits. All parameters involved are fully investigated in the scope of the validation work.

Accurate results from amino acid analysis require purified protein and peptide samples. Buffer components (e.g., salts, urea, detergents) can interfere with the amino acid analysis and are removed from the sample before analysis. Methods that utilize postcolumn derivatization of the amino acids are generally not affected by buffer components to the extent seen with precolumn derivatization methods. It is desirable to limit the number of sample manipulations to reduce potential background contamination, to improve analyte recovery, and to reduce labor. Common techniques used to remove buffer components from protein samples include the following methods: (1) injecting the protein sample onto a reverse-phase HPLC system, removing the protein with a volatile solvent containing a sufficient organic component, and drying the sample in a vacuum centrifuge; (2) dialysis against a volatile buffer or water; (3) centrifugal ultrafiltration for buffer replacement with a volatile buffer or water; (4) precipitating the protein from the buffer using an organic solvent (e.g., acetone); and (5) gel filtration.

It is recommended that an internal standard be used to monitor physical and chemical losses and variations during amino acid analysis. An accurately known amount of internal standard can be added to a protein solution prior to hydrolysis. The recovery of the internal standard gives the general recovery of the amino acids from the protein solution. Free amino acids, however, do not behave in the same way as protein-bound amino acids during hydrolysis because their rates of release or destruction are variable. Therefore, the use of an internal standard to correct for losses during hydrolysis may give unreliable results. It will be necessary to take this particular point into consideration when interpreting the results. Internal standards can also be added to the mixture of amino acids after hydrolysis to correct for differences in sample application and changes in reagent stability and flow rates. Ideally, an internal standard is an unnaturally occurring primary amino acid that is commercially available and inexpensive. It should also be stable during hydrolysis, its response factor should be linear with concentration, and it needs to elute with a unique retention time without overlapping other amino acids. Commonly used amino acid standards include norleucine, nitrotyrosine, and -aminobutyric acid.

Hydrolysis of protein and peptide samples is necessary for amino acid analysis of these molecules. The glassware used for hydrolysis must be very clean to avoid erroneous results. Glove powders and fingerprints on hydrolysis tubes may cause contamination. To clean glass hydrolysis tubes, boil tubes for 1 hour in 1 N hydrochloric acid or soak tubes in concentrated nitric acid or in a mixture of concentrated hydrochloric acid and concentrated nitric acid (1:1). Clean hydrolysis tubes are rinsed with high-purity water followed by a rinse with HPLC grade methanol, dried overnight in an oven, and stored covered until use. Alternatively, pyrolysis of clean glassware at 500 for 4 hours may also be used to eliminate contamination from hydrolysis tubes. Adequate disposable laboratory material can also be used.

Acid hydrolysis is the most common method for hydrolyzing a protein sample before amino acid analysis. The acid hydrolysis technique can contribute to the variation of the analysis due to complete or partial destruction of several amino acids. Tryptophan is destroyed; serine and threonine are partially destroyed; methionine might undergo oxidation; and cysteine is typically recovered as cystine (but cystine recovery is usually poor because of partial destruction or reduction to cysteine). Application of adequate vacuum (less than 200 µm of mercury or 26.7 Pa) or introduction of an inert gas (argon) in the headspace of the reaction vessel can reduce the level of oxidative destruction. In peptide bonds involving isoleucine and valine, the amido bonds of Ile-Ile, Val-Val, Ile-Val, and Val-Ile are partially cleaved; and asparagine and glutamine are deamidated, resulting in aspartic acid and glutamic acid, respectively. The loss of tryptophan, asparagine, and glutamine during an acid hydrolysis limits quantitation to 17 amino acids. Some of the hydrolysis techniques described are used to address these concerns. Some of the hydrolysis techniques described (i.e., Methods 4–11) may cause modifications to other amino acids. Therefore, the benefits of using a given hydrolysis technique are weighed against the concerns with the technique and are tested adequately before employing a method other than acid hydrolysis.

A time-course study (i.e., amino acid analysis at acid hydrolysis times of 24, 48, and 72 hours) is often employed to analyze the starting concentration of amino acids that are partially destroyed or slow to cleave. By plotting the observed concentration of labile amino acids (i.e., serine and threonine) versus hydrolysis time, the line can be extrapolated to the origin to determine the starting concentration of these amino acids. Time-course hydrolysis studies are also used with amino acids that are slow to cleave (e.g., isoleucine and valine). During the hydrolysis time course, the analyst will observe a plateau in these residues. The level of this plateau is taken as the residue concentration. If the hydrolysis time is too long, the residue concentration of the sample will begin to decrease, indicating destruction by the hydrolysis conditions.

An acceptable alternative to the time-course study is to subject an amino acid calibration standard to the same hydrolysis conditions as the test sample. The amino acid in free form may not completely represent the rate of destruction of labile amino acids within a peptide or protein during the hydrolysis. This is especially true for peptide bonds that are slow to cleave (e.g., Ile-Val bonds). However, this technique will allow the analyst to account for some residue destruction. Microwave acid hydrolysis has been used and is rapid but it requires special equipment as well as special precautions. The optimal conditions for microwave hydrolysis must be investigated for each individual protein/peptide sample. The microwave hydrolysis technique typically requires only a few minutes, but even a deviation of 1 minute may give inadequate results (e.g., incomplete hydrolysis or destruction of labile amino acids). Complete proteolysis, using a mixture of proteases, has been used but can be complicated, requires the proper controls, and is typically more applicable to peptides than proteins. [NOTE—During initial analyses of an unknown protein, experiments with various hydrolysis time and temperature conditions are conducted to determine the optimal conditions.]

METHOD 1

Hydrolysis Solution: 6 N hydrochloric acid containing 0.1% to 1.0% of phenol.

Procedure—

METHOD 2

Hydrolysis Solution: 2.5 M MESA solution.

Vapor Phase Hydrolysis— About 1 to 100 µg of the protein/peptide under test is dried in a hydrolysis tube. The hydrolysis tube is placed in a larger tube with about 200 µL of the Hydrolysis Solution. The larger tube is sealed in vacuum (about 50 µm of mercury or 6.7 Pa) to vaporize the Hydrolysis Solution. The hydrolysis tube is heated to between 170 to 185 for about 12.5 minutes. After hydrolysis, the hydrolysis tube is dried in vacuum for 15 minutes to remove the residual acid.

METHOD 3

Hydrolysis Solution: a solution containing 7 M hydrochloric acid, 10% of trifluoroacetic acid, 20% of thioglycolic acid, and 1% of phenol.

Vapor Phase Hydrolysis— About 10 to 50 µg of the protein/peptide under test is dried in a sample tube. The sample tube is placed in a larger tube with about 200 µL of the Hydrolysis Solution. The larger tube is sealed in vacuum (about 50 µm of mercury or 6.7 Pa) to vaporize the TGA. The sample tube is heated to 166 for about 15 to 30 minutes. After hydrolysis, the sample tube is dried in vacuum for 5 minutes to remove the residual acid. Recovery of tryptophan by this method may be dependent on the amount of sample present.

METHOD 4

Oxidation Solution— The performic acid is prepared fresh by mixing formic acid and 30 percent hydrogen peroxide (9:1), and incubated at room temperature for 1 hour.

Procedure— The protein/peptide sample is dissolved in 20 µL of formic acid, and heated at 50 for 5 minutes; then 100 µL of the Oxidation Solution is added. In this reaction, cysteine is converted to cysteic acid and methionine is converted to methionine sulfone. The oxidation is allowed to proceed for 10 to 30 minutes. The excess reagent is removed from the sample in a vacuum centrifuge. This technique may cause modifications to tyrosine residues in the presence of halides. The oxidized protein can then be acid hydrolyzed using Method 1 or Method 2.

METHOD 5

Hydrolysis Solution: 6 N hydrochloric acid containing 0.2% of phenol, to which sodium azide is added to obtain a final concentration of 0.2% (w/v). The added phenol prevents halogenation of tyrosine.

Liquid Phase Hydrolysis— The protein/peptide hydrolysis is conducted at about 110 for 24 hours. During the hydrolysis, the cysteine-cystine present in the sample is converted to cysteic acid by the sodium azide present in the Hydrolysis Solution. This technique allows better tyrosine recovery than Method 4, but it is not quantitative for methionine. Methionine is converted to a mixture of the parent methionine and its two oxidative products, methionine sulfoxide and methionine sulfone.

METHOD 6

Hydrolysis Solution: 6 N hydrochloric acid containing 0.1% to 1.0% of phenol, to which DMSO is added to obtain a final concentration of 2% (v/v).

Vapor Phase Hydrolysis— The protein/peptide hydrolysis is conducted at about 110 for 24 hours. During the hydrolysis, the cysteine-cystine present in the sample is converted to cysteic acid by the DMSO present in the Hydrolysis Solution. As an approach to limit variability and compensate for partial destruction, it is recommended to evaluate the cysteic acid recovery from oxidative hydrolyses of standard proteins containing 1 to 8 mol of cysteine. The response factors from protein/peptide hydrolysates are typically about 30% lower than those for nonhydrolyzed cysteic acid standards. Because histidine, methionine, tyrosine, and tryptophan are also modified, a complete compositional analysis is not obtained with this technique.

METHOD 7

Reducing Solution— Transfer 83.3 µL of pyridine, 16.7 µL of 4-vinylpyridine, 16.7 µL of tributylphosphine, and 83.3 µL of water to a suitable container, and mix.

Procedure— Add the protein/peptide (between 1 and 100 µg) to a hydrolysis tube, and place in a larger tube. Transfer the Reducing Solution to the large tube, seal in vacuum (about 50 µm of mercury or 6.7 Pa), and incubate at about 100 for 5 minutes. Then remove the inner hydrolysis tube, and dry it in a vacuum desiccator for 15 minutes to remove residual reagents. The pyridylethylated protein/peptide can then be acid hydrolyzed using previously described procedures. The pyridylethylation reaction is performed simultaneously with a protein standard sample containing 1 to 8 mol of cysteine to improve accuracy in the pyridylethyl-cysteine recovery. Longer incubation times for the pyridylethylation reaction can cause modifications to the -amino terminal group and the -amino group of lysine in the protein.

METHOD 8

Stock Solutions— Prepare and filter three solutions: 1 M Tris hydrochloride (pH 8.5) containing 4 mM edetate disodium (Stock Solution 1), 8 M guanidine hydrochloride (Stock Solution 2), and 10% of 2-mercaptoethanol in water (Stock Solution 3).

Reducing Solution— Prepare a mixture of Stock Solution 2 and Stock Solution 1 (3:1) to obtain a buffered solution of 6 M guanidine hydrochloride in 0.25 M Tris hydrochloride.

Procedure— Dissolve about 10 µg of the test sample in 50 µL of the Reducing Solution, and add about 2.5 µL of Stock Solution 3. Store under nitrogen or argon for 2 hours at room temperature in the dark. To achieve the pyridylethylation reaction, add about 2 µL of 4-vinylpyridine to the protein solution, and incubate for an additional 2 hours at room temperature in the dark. The protein/peptide is desalted by collecting the protein/peptide fraction from a reverse-phase HPLC separation. The collected sample can be dried in a vacuum centrifuge before acid hydrolysis.

METHOD 9

Stock Solutions— Prepare as directed for Method 8.

Carboxymethylation Solution— Prepare a solution containing 100 mg of iodoacetamide per mL of alcohol.

Buffer Solution— Use the Reducing Solution, prepared as directed for Method 8.

Procedure— Dissolve the test sample in 50 µL of the Buffer Solution, and add about 2.5 µL of Stock Solution 3. Store under nitrogen or argon for 2 hours at room temperature in the dark. Add the Carboxymethylation Solution in a 1.5 fold ratio per total theoretical content of thiols, and incubate for an additional 30 minutes at room temperature in the dark. [NOTE—If the thiol content of the protein is unknown, then add 5 µL of 100 mM iodoacetamide for every 20 nmol of protein present.] The reaction is stopped by adding excess of 2-mercaptoethanol. The protein/peptide is desalted by collecting the protein/peptide fraction from a reverse-phase HPLC separation. The collected sample can be dried in a vacuum centrifuge before acid hydrolysis. The S-carboxyamidomethylcysteine formed will be converted to S-carboxymethyl-cysteine during acid hydrolysis.

METHOD 10

Reducing Solution: a solution containing 10 mg of dithiodiglycolic acid (or dithiodipropionic acid) per mL of 0.2 M sodium hydroxide.

Procedure— Transfer about 20 µg of the test sample to a hydrolysis tube, and add 5 µL of the Reducing Solution. Add 10 µL of isopropyl alcohol, and then remove all of the sample liquid by vacuum centrifugation. The sample is then hydrolyzed using Method 1. This method has the advantage that other amino acid residues are not derivatized by side reactions, and the sample does not need to be desalted prior to hydrolysis.

METHOD 11

Reducing Solutions— Prepare and filter three solutions: a solution of 10 mM trifluoroacetic acid (Solution 1), a solution of 5 M guanidine hydrochloride and 10 mM trifluoroacetic acid (Solution 2), and a freshly prepared solution of dimethylformamide containing 36 mg of BTI per mL (Solution 3).

Procedure— In a clean hydrolysis tube, transfer about 200 µg of the test sample, and add 2 mL of Solution 1 or Solution 2 and 2 mL of Solution 3. Seal the hydrolysis tube in vacuum. Heat the sample at 60 for 4 hours in the dark. The sample is then dialyzed with water to remove the excess reagents. Extract the dialyzed sample three times with equal volumes of n-butyl acetate, and then lyophilize. The protein can then be acid hydrolyzed using previously described procedures. The -, -diaminopropionic and -, -diaminobutyric acid residues do not typically resolve from the lysine residues upon ion-exchange chromatography based on amino acid analysis. Therefore, when using ion-exchange as the mode of amino acid separation, the asparagine and glutamine contents are the quantitative difference in the aspartic acid and glutamic acid assayed contents with un-derivatized and BTI-derivatized acid hydrolysis. [NOTE—The threonine, methionine, cysteine, tyrosine, and histidine assayed content can be altered by BTI derivatization; a hydrolysis without BTI will have to be performed if the analyst is interested in the protein/peptide content of these other amino acid residues.]

Many amino acid analysis techniques exist, and the choice of any one technique often depends on the sensitivity required from the assay. In general, about one-half of the amino acid analysis techniques employed rely on the separation of the free amino acids by ion-exchange chromatography followed by postcolumn derivatization (e.g., with ninhydrin or o-phthalaldehyde). Postcolumn detection techniques can be used with samples that contain small amounts of buffer components, such as salts and urea, and generally require between 5 and 10 µg of protein sample per analysis. The remaining amino acid techniques typically involve precolumn derivatization of the free amino acids (e.g., phenyl isothiocyanate; 6-aminoquinolyl-N-hydroxysuccinimidyl carbonate; (dimethylamino)azobenzenesulfonyl chloride; 9-fluorenyl-methylchloroformate; and 7-fluoro-4-nitrobenzo-2-oxa-1,3-diazole) followed by reverse-phase HPLC. Precolumn derivatization techniques are very sensitive and usually require between 0.5 and 1.0 µg of protein sample per analysis but may be influenced by buffer salts in the samples. Precolumn derivatization techniques may also result in multiple derivatives of a given amino acid, which complicates the result interpretation. Postcolumn derivatization techniques are generally influenced less by performance variation of the assay than precolumn derivatization techniques.

The following Methods may be used for quantitative amino acid analysis. Instruments and reagents for these procedures are available commercially. Furthermore, many modifications of these methodologies exist with different reagent preparations, reaction procedures, chromatographic systems, etc. Specific parameters may vary according to the exact equipment and procedure used. Many laboratories will utilize more than one amino acid analysis technique to exploit the advantages offered by each. In each of these Methods, the analog signal is visualized by means of a data acquisition system, and the peak areas are integrated for quantification purposes.

METHOD 1—POSTCOLUMN NINHYDRIN DETECTION

Mobile Phase Preparation—

Postcolumn Reagent— Transfer about 18 g of ninhydrin and 0.7 g of hydrindantin to 900 mL of a solution containing 76.7% of dimethyl sulfoxide, 0.7% of dihydrate lithium acetate, and 0.1% of acetic acid, and mix for at least 3 hours under inert gas, such as nitrogen. [NOTE—This reagent is stable for 30 days if kept between 2 and 8 under inert gas.]

Buffer Solution— Prepare a solution containing 2% of anhydrous sodium citrate, 1% of hydrochloric acid, 0.5% of thiodiglycol, and 0.1% of benzoic acid in water, and adjust to a pH of 2.

Chromatographic System— The liquid chromatograph is equipped with a detector with appropriate interference filters at 440, 570, or 690 nm and a 4.0-mm × 120-mm column that contains 7.5-µm sulfonated styrene-divinylbenzene copolymer packing. The flow rate is about 14 mL per hour. The system is programmed as follows. Initially equilibrate the column with Solution A; at 25 minutes, the composition of the Mobile Phase is changed to 100% Solution B; and at 37 minutes, the composition is changed to 100% Solution C. At 75 minutes into the run, the last amino acid has been eluted from the column, and the column is regenerated with Column Regeneration Solution for 1 minute. The column is then equilibrated with Solution A for 11 minutes before the next injection. The column temperature is programmed as follows. The initial temperature is 48; after 11.5 minutes, the temperature is increased to 65 at a rate of 3 per minute; at about 35 minutes, the temperature is increased to 77 at a rate of 3 per minute; and finally at about 52 minutes, the temperature is decreased to 48 at a rate of 3 per minute.

Procedure and Postcolumn Reaction— Reconstitute the lyophilized protein/peptide hydrolysate in the Buffer Solution, inject an appropriate amount into the chromatograph, and proceed as directed for Chromatographic System. As the amino acids are eluted from the column, they are mixed with the Postcolumn Reagent, which is delivered at a flow rate of 7 mL per hour, through a tee. After mixing, the column effluent and the Postcolumn Reagent pass through a tubular reactor at a temperature of 135, where a characteristic purple or yellow color is developed. From the reactor, the liquid passes through a colorimeter with a 12-mm flow-through cuvette. The light emerging from the cuvette is split into three beams for analysis by the detector with interference filters at 440, 570, or 690 nm. The 690-nm signal may be electronically subtracted from the other signals for improved signal-to-noise ratios. The 440-nm (imino acids) and the 570-nm (amino acids) signals may be added in order to simplify data handling.

METHOD 2—POSTCOLUMN OPA FLUOROMETRIC DERIVATIZATION

Mobile Phase Preparation—

Postcolumn Reagent Preparation—

Chromatographic System— The liquid chromatograph is equipped with a fluorometric detector set to an excitation wavelength of 348 nm and an emission wavelength of 450 nm and a 4.0-mm × 150-mm column that contains 7.5-µm packing L17. The flow rate is about 0.3 mL per minute, and the column temperature is set at 50. The system is programmed as follows. The column is equilibrated with Solution A; over the next 20 minutes, the composition of the Mobile Phase is changed linearly to 85% Solution A and 15% Solution B; then there is a step change to 40% Solution A and 60% Solution B; over the next 18 minutes, the composition is changed linearly to 100% Solution B and held for 7 minutes; then there is a step change to 100% Solution C, and this is held for 6 minutes; then there is a step change to Solution A, and this composition is maintained for the next 8 minutes.

Procedure and Postcolumn Reaction— Inject about 1.0 nmol of each amino acid under test into the chromatograph, and proceed as directed for Chromatographic System. As the effluent leaves the column, it is mixed with the Hypochlorite Reagent. The mixture passes through the first postcolumn reactor which consists of stainless steel 0.5-mm × 2-m tubing. A second postcolumn reactor of similar design is placed immediately downstream from the first postcolumn reactor and is used for the OPA postcolumn reaction. The flow rates for both the Hypochlorite Reagent and the OPA Reagent are 0.2 mL per minute, resulting in a total flow rate (i.e., Hypochlorite Reagent, OPA Reagent, and column effluent) of 0.7 mL per minute exiting from the postcolumn reactors. Postcolumn reactions are conducted at 55. This results in a residence time of about 33 seconds in the OPA postcolumn reactor. After postcolumn derivatization, the column effluent passes through the fluorometric detector.

METHOD 3—PRECOLUMN DETERMINATION

Mobile Phase Preparation—

Derivatization Reagent Preparation—

Sample Derivatization Procedure— Dissolve the lyophilized test sample in 100 µL of the Coupling Buffer, and then dry in a vacuum centrifuge to remove any hydrochloride if a protein hydrolysis step was used. Dissolve the test sample in 100 µL of Coupling Buffer, add 5 µL of PITC, and incubate at room temperature for 5 minutes. The test sample is again dried in a vacuum centrifuge, and is dissolved in 250 µL of Sample Solvent.

Chromatographic System— The liquid chromatograph is equipped with a 254-nm detector and a 4.6-mm × 250-mm column that contains 5-µm packing L1. The flow rate is about 1 mL per minute, and the column temperature is maintained at 52. The system is programmed as follows. The column is equilibrated with Solution A; over the next 15 minutes, the composition of the Mobile Phase is changed linearly to 85% Solution A and 15% Solution B; over the next 15 minutes, the composition is changed linearly to 50% Solution A and 50% Solution B; then there is a step change to 100% Solution C, and this is held for 10 minutes; then there is a step change to 100% Solution A, and the column is allowed to equilibrate before the next injection.

Procedure— Inject about 1.0 nmol of each PITC-amino acid under test (10-µL sample in Sample Solvent) into the chromatograph, and proceed as directed for Chromatographic System.

METHOD 4—PRECOLUMN AQC DERIVATIZATION

Mobile Phase Preparation—

Sample Derivatization Procedure— Dissolve about 2 µg of the test sample in 20 µL of 15 mM hydrochloric acid, and dilute with 0.2 M borate buffer (pH 8.8) to 80 µL. The derivatization is initiated by the addition of 20 µL of 10 mM AQC in acetonitrile, and allowed to proceed for 10 minutes at room temperature.

Chromatographic System— The liquid chromatograph is equipped with a fluorometric detector set at an excitation wavelength of 250 nm and an emission wavelength of 395 nm and a 3.9-mm × 150-mm column that contains 4-µm packing L1. The flow rate is about 1 mL per minute, and the column temperature is maintained at 37. The system is programmed as follows. The column is equilibrated with Solution A; over the next 0.5 minute, the composition of the Mobile Phase is changed linearly to 98% Solution A and 2% Solution B; then over the next 14.5 minutes to 93% Solution A and 7% Solution B; then over the next 4 minutes to 87% Solution A and 13% Solution B; over the next 14 minutes to 68% Solution A and 32% Solution B; then there is a step change to 100% Solution B for a 5-minute wash; over the next 10 minutes, there is a step change to 100% Solution A; and the column is allowed to equilibrate before the next injection.

Procedure— Inject about 0.05 nmol of each AQC-amino acid under test into the chromatograph, and proceed as directed for Chromatographic System.

METHOD 5—PRECOLUMN OPA DERIVATIZATION

Mobile Phase Preparation—

Derivatization Reagent— Dissolve 50 mg of OPA in 1.25 mL of methanol (protein sequencing grade). Add 50 µL of 2-mercaptoethanol and 11.2 mL of 0.4 M sodium borate (pH 9.5), and mix. [NOTE—This reagent is stable for 1 week.]

Sample Derivatization Procedure— Transfer about 5 µL of the test sample to an appropriate container, add 5 µL of the Derivatization Reagent, and mix. After 1 minute, add not less than 20 µL of 0.1 M sodium acetate (pH 7.0). Use 20 µL of this solution for analysis. [NOTE—Use of an internal standard (e.g., norleucine) is recommended for quantitative analysis because of potential reagent volume variations in the sample derivatization. The sample derivatization is performed in an automated on-line fashion. Because of the instability of the OPA-amino acid derivative, HPLC separation and analysis are performed immediately following derivatization.]

Chromatographic System— The liquid chromatograph is equipped with a fluorometric detector set at an excitation wavelength of 348 nm and an emission wavelength of 450 nm and a 4.6-mm × 75-mm column that contains 3-µm packing L3. The flow rate is about 1.7 mL per minute, and the column temperature is maintained at 37. The system is programmed as follows. The column is equilibrated with 92% Solution A and 8% Solution B; over the next 2 minutes, the composition of the Mobile Phase is changed to 83% Solution A and 17% Solution B, and held for an additional 3 minutes; then changed to 54% Solution A and 46% Solution B over the next 5 minutes, and held for an additional 2 minutes; then changed to 34% Solution A and 66% Solution B over the next 2 minutes, and held for 1 minute; then over the next 0.3 minute changed to 20% Solution A and 80% Solution B, and held for an additional 2.6 minutes; and then finally over 0.6 minute changed to 92% Solution A and 8% Solution B, and held for an additional 0.6 minute.

Procedure— Inject about 0.02 nmol of each OPA-amino acid under test into the chromatograph, and proceed as directed for Chromatographic System.

METHOD 6—POSTCOLUMN DABS-Cl DERIVATIZATION

Mobile Phase Preparation—

Derivatization Reagent Preparation—

Sample Derivatization Procedure— Dissolve the test sample in 20 µL of Sample Buffer, add 40 µL of Derivatization Reagent, and mix. The sample container is sealed with a silicon-rubber stopper, and heated to 70 for 10 minutes. During the sample heating, the mixture will become completely soluble. After the derivatization, dilute the test sample with an appropriate quantity of the Sample Dilution Buffer.

Chromatographic System— The liquid chromatograph is equipped with a 436-nm detector and a 4.6-mm × 250-mm column that contains packing L1. The flow rate is about 1 mL per minute, and the column temperature is maintained at 40. The system is programmed as follows. The column is equilibrated with 85% Solution A and 15% Solution B; over the next 20 minutes, the composition of the Mobile Phase is changed to 60% Solution A and 40% Solution B; over the next 12 minutes, the composition is changed to 30% Solution A and 70% Solution B, and held for an additional 2 minutes.

Procedure— Inject about 0.05 nmol of the DABS-amino acids into the chromatograph, and proceed as directed for Chromatographic System.

METHOD 7—PRECOLUMN FMOC-Cl DERIVATIZATION

Mobile Phase Preparation—

Derivatization Reagent Preparation—

Sample Derivatization Procedure— To 0.4 mL of the test sample add 0.1 mL of Borate Buffer and 0.5 mL of FMOC-Cl Reagent. After about 40 seconds, extract the mixture with 2 mL of pentane, and then extract again with fresh pentane. The aqueous solution with amino acid derivatives is then ready for injection.

Chromatographic System— The liquid chromatograph is equipped with a fluorometric detector set at an excitation wavelength of 260 nm and an emission wavelength of 313 nm and a 4.6-mm × 125-mm column that contains 3-µm packing L1. The flow rate is about 1.3 mL per minute. The system is programmed as follows. The column is equilibrated with Solution A, and this composition is maintained for 3 minutes; over the next 9 minutes, it is changed to 100% Solution B; then over the next 0.5 minute, the flow rate is increased to 2 mL per minute, and held until the final FMOC-amino acid is eluted from the column. The total run time is about 20 minutes.

Procedure— Inject not less than 0.01 nmol of each FMOC-amino acid under test into the chromatograph, and proceed as directed for Chromatographic System. The FMOC-histidine derivative will generally give a lower response than the other derivatives.

METHOD 8—PRECOLUMN NBD-F DERIVATIZATION

Mobile Phase Preparation—

Derivatization Reagent Preparation—

Sample Derivatization Procedure— Dissolve the test sample in 20 µL of Sample buffer, add 10 µL of Derivatization Reagent, and mix. The sample container is heated at 60 for 5 minutes. After the derivatization, dilute the test sample with 300 µL of Solution A.

Chromatographic System— The liquid chromatograph is equipped with a fluorometric detector set at an excitation wavelength of 480 nm and an emission wavelength of 530 nm and a 4.6-mm × 150-mm column that contains 5-µm particle size ODS silica packing. The flow rate is about 1.0 mL per minute, and the column temperature is maintained at 40. The system is programmed as follows. The column is equilibrated with 94% Solution A and 6% Solution B; over the next 16 minutes, the composition is changed linearly to 63% Solution A and 37% Solution B; over the next 5 minutes, the composition is changed linearly to 62% Solution A and 38% Solution B; over the next 9 minutes, the composition is changed linearly to 100% Solution B, and held for an additional 5 minutes; then finally over 2 minutes, the composition is changed linearly to 94% Solution A and 6% Solution B; and then the column is allowed to equilibrate before the next injection.

Procedure— Inject about 15 pmol of each NBD-amino acid under test into the chromatograph, and proceed as directed for Chromatographic System.

When determining the amino acid content of a protein/peptide hydrolysate, it should be noted that the acid hydrolysis step destroys tryptophan and cysteine. Serine and threonine are partially destroyed by acid hydrolysis, while isoleucine and valine residues may be only partially cleaved. Methionine can undergo oxidation during acid hydrolysis, and some amino acids (e.g., glycine and serine) are common contaminants. Application of adequate vacuum (less than 200 um µm of mercury or 26.7 Pa) or introduction of inert gas (argon) in the headspace of the reaction vessel during vapor phase hydrolysis can reduce the level of oxidative destruction. Therefore, the quantitative results obtained for cysteine, tryptophan, threonine, isoleucine, valine, methionine, glycine, and serine from a protein/peptide hydrolysate may be variable and may warrant further investigation and consideration.

CALCULATIONS

Amino Acid Mole Percent— This is the number of specific amino acid residues per 100 residues in a protein. This result may be useful for evaluating amino acid analysis data when the molecular weight of the protein/peptide under investigation is unknown. This information can be used to corroborate the identity of a protein and has other applications. Carefully identify and integrate the peaks obtained as directed for each Procedure. Calculate the mole percent for each amino acid present in the test sample by the formula: in which rU is the peak response, in nmol, of the amino acid under test; and r is the sum of peak responses, in nmol, for all amino acids present in the test sample. Comparison of the mole percent of the amino acids under test to data from known proteins can help establish or corroborate the identity of the sample protein.

Unknown Protein Samples— This data analysis technique can be used to estimate the protein concentration of an unknown protein sample using the amino acid analysis data. Calculate the mass, in µg, of each recovered amino acid by the formula: in which m is the recovered quantity, in nmol, of the amino acid under test; and MW is the average molecular weight, in mg, for that amino acid, corrected for the weight of the water molecule that was eliminated during peptide bond formation. The sum of the masses of the recovered amino acids will give an estimate of the total mass of the protein analyzed after appropriate correction for partially and completely destroyed amino acids. If the molecular weight of the unknown protein is available (i.e., by SDS-PAGE analysis or mass spectroscopy), the amino acid composition of the unknown protein can be predicted. Calculate the number of residues of each amino acid by the formula: in which m is the recovered quantity, in nmol, of the amino acid under test; M is the total mass, in µg, of the protein; and MWT is the molecular weight, in mg, of the unknown protein.

Known Protein Samples— This data analysis technique can be used to investigate the amino acid composition and protein concentration of a protein sample of known molecular weight and amino acid composition using the amino acid analysis data. When the composition of the protein being analyzed is known, one can exploit the fact that some amino acids are recovered well, while other amino acid recoveries may be compromised because of complete or partial destruction (e.g., tryptophan, cysteine, threonine, serine, methionine), incomplete bond cleavage (i.e., for isoleucine and valine), and free amino acid contamination (i.e., by glycine and serine).

The migration velocity of the analyte under an electric field of intensity, E, is determined by the electrophoretic mobility of the analyte and the electroosmotic mobility of the buffer inside the capillary. The electrophoretic mobility of a solute (µep) depends on the characteristics of the solute (electrical charge, molecular size, and shape) and the characteristics of the buffer in which the migration takes place (type and ionic strength of the electrolyte, pH, viscosity, and additives). The electrophoretic velocity (Vep) of a solute, assuming a spherical shape, is as follows:

in which q is the effective charge of the particle; is the viscosity of the buffer; r is the size of the solute ion; V is the applied voltage; and L is the total length of the capillary.

When an electric field is applied through the capillary filled with buffer, a flow of solvent is generated inside the capillary called electroosmotic flow. Its velocity depends on the electroosmotic mobility (µeo) which in turn depends on the charge density on the capillary internal wall and the buffer characteristics. The electroosmotic velocity (Veo) is as follows:

in which is the dielectric constant of the buffer; is the zeta potential of the capillary surface; and the other terms are as defined above.

The electrophoretic and electroosmotic mobilities of the analyte may act in the same direction or in opposite directions, depending on the charge (positive or negative) of the solute, and the velocity of the solute (v) is as follows:

The sum or the difference between the two velocities (Vep and Veo) is used depending on whether the mobilities act in the same or opposite directions. Under conditions with a fast Veo, with respect to the Vep of the solutes, both negative and positive charged analytes can be separated in the same run. The time (t) taken by the solute to migrate the distance (l) from the injection end of the capillary to the detection point (capillary effective length) is as follows:

in which the other terms are as defined above.

In general, the fused-silica capillaries used in electrophoresis bear negative charges on the inner wall, producing electroosmotic flow towards the cathode. The electroosmotic flow has to remain constant from run to run to obtain good reproducibility in the migration velocity of the solutes. For some applications, it might be necessary to reduce or suppress the electroosmotic flow by modifying the inner wall of the capillary or by changing the pH of the buffer solution.

When the sample is introduced in the capillary, each analyte ion of the sample migrates within the background electrolyte as an independent zone according to its electrophoretic mobility. The spreading of each solute band (zone dispersion) results from a different phenomena. Under ideal conditions, the sole contribution to the solute-zone broadening is molecular diffusion of the solute along the capillary (longitudinal diffusion). In this case, the efficiency of the zone is expressed as the number of theoretical plates (N), as follows:

in which D is the molecular diffusion of the solute in the buffer; and the other terms are as defined above.

From a practical point of view, other phenomena such as heat dissipation, sample adsorption onto the capillary wall, mismatched conductivity between sample and buffer, length of the injection plug, detector cell size, and unleveled buffer reservoirs can also significantly contribute to band dispersion. Separation between two bands (expressed by the resolution, RS) can be achieved by modification of the electrophoretic mobility of the analytes, by the electroosmotic mobility induced by capillary, and by increasing the efficiency for the band of each analyte as follows:

in which µepa and µepb are the electrophoretic mobilities of the two compounds to be separated; bar(µ)ep is the average electrophoretic mobility of the two solutes calculated as:

and the other terms are as defined above.

An apparatus for capillary electrophoresis is composed of a high voltage controllable power supply; two buffer reservoirs held at the same level and containing specified anodic and cathodic solutions; two electrodes assemblies (cathode and anode) immersed in the buffer reservoirs and connected to the power supply; a separation capillary usually made of fused-silica, with sometimes an optical viewing window aligned with detector, depending on the detector, with the ends of the capillary placed in the buffer reservoirs and the capillary being filled with a solution specified in a given monograph; a suitable injection system; a detector capable of monitoring the amount of substance of interest passing through a segment of the separation capillary at a given time, generally based on absorption spectrophotometry (UV and visible), fluorimetry, conductimetric, amperometric, or mass spectrometric detection, depending on the specific applications, or even indirect detection to detect non-UV-absorbing and nonfluorescent compounds; and a thermostatic system capable of maintaining the temperature inside the capillary.

The method of injection of samples and its automation is critical for precise quantitative analysis. Methods of injection include gravity, pressure or vacuum, or electrokinetic injection. The amount of each sample component introduced electrokinetically depends on its electrophoretic mobility, thus possibly biasing the results.

It is expected that the capillary, the buffer solutions, the preconditioning method, the sample solution, and the migration conditions will be specified in the individual monograph. The electrolytic solution employed may be filtered to remove particles and degassed to avoid bubble formation that could interfere with the detection system. To achieve reproducible migration time of the solutes, if would be necessary to develop, for each analytical method, a rigorous rinsing routine after each injection.

In free solution capillary electrophoresis, analytes are separated in a capillary containing only buffer without any anticonvective medium. In this technique, separation takes place because the different components of the sample migrate as discrete bands with different velocities. The velocity of each band depends on the electrophoretic mobility of the solute and the electroosmotic flow on the capillary. Coated capillaries, with reduced electroosmotic flow, can be used to increase the separation capacity of those substances absorbing on fused silica surfaces.

This mode of capillary electrophoresis is appropriate for the analysis of small (MW < 2000) and large molecules (2000 < MW < 100,000). Due to the high efficiency achieved, molecules having only minute differences in their charge-to-mass ratio can be separated. This method also allows the separation of chiral compounds by adding chiral selectors to the separation buffer. The optimization of the separations requires consideration of a number of instrumental and electrolytic solution parameters.

INSTRUMENTAL PARAMETERS

Voltage— The separation time is universally proportional to applied voltage. However, an increase in the voltage used can cause excessive heat production, giving rise to temperature and viscosity gradients in the buffer inside the capillary, which causes band broadening and decreases resolution.

Temperature— The main effect of temperature is observed on buffer viscosity and electrical conductivity, thus affecting migration velocity. In some cases, an increase in capillary temperature can cause a conformational change of some proteins, modifying their migration time and the efficiency of the separation.

Capillary— The length and internal diameter of the capillary affects the analysis time, the efficiency of separations, and the load capacity. Increasing both effective length and total length can decrease the electric fields, at a constant voltage, which will increase migration time. For a given buffer and electric field, heat dissipation (thus sample band broadening) depends on the internal diameter of the capillary. The latter also affects the detection limit, depending on the sample volume injected into the capillary and the detection system used.

ELECTROLYTIC SOLUTION PARAMETERS

Buffer Type and Concentrations— Suitable buffers for capillary electrophoresis have an appropriate buffer capacity in the pH range of choice and low mobility to minimize current generation.

Buffer pH— The pH of the buffer can affect separation by modifying the charge of the analyte or other additives and by changing the electroosmotic flow. For protein and peptide separation, a change in the pH of the buffer from above the isoelectric point to below the isoelectric point changes the net charge of the solute from negative to positive. An increase in the buffer pH generally increases the electroosmotic flow.

Organic Solvents— Organic modifiers, such as methanol, acetonitrile, and others, are added to the aqueous buffer to increase the solubility of the solute or other additives and/or to affect the ionization degree of the sample components. The addition of these organic modifiers to the buffer generally causes a decrease in the electroosmotic flow.

Additives for Chiral Separations— To separate optical isomers, a chiral selector is added to the separation buffer. The most commonly used chiral selectors are cyclodextrins, although in some cases crown ethers, certain polysaccharides, or even proteins can be used. Because chiral recognition is governed by the different interactions between the chiral selector and each of the enantiomers, the resolution achieved for the chiral compounds depends largely on the type of chiral selector used. While developing a given separation it may be useful to test cyclodextrins having a different cavity size (-, -, or G-cyclodextrin) or modified cyclodextrins with neutral (methyl, ethyl, hydroxyalkyl, etc.) or ionizable (aminomethyl, carboxymethyl, sulfobutylether, etc.) moities. The resolution of chiral separations is also controlled by the concentration of the chiral selector, the composition and pH of the buffer, and the separation temperature. Organic additives, such as methanol or urea, can also affect the resolution of separation.

Separation takes place inside a capillary filled with a polymer acting as a molecular sieve. The smaller components in the sample move faster along the capillary than the larger ones. This method can be used for separation of biopolymers-proteins, and DNA fragments, according to their molecular mass.

CHARACTERISTICS OF CHEMICAL and PHYSICAL GELS

Chemical Gels— Chemical gels are prepared inside the capillary by reaction of monomers. One example of such a gel is a cross-linked polyacrylamide. This type of gel is bonded to the fused-silica wall and cannot be removed without destroying the capillary. For protein analysis, the separation buffer usually contains sodium dodecyl sulfate and the sample is denatured by heating in a mixture of sodium dodecyl sulfate and 2-mercaptoethanol or dithiothreitol before injection. Optimization of separation in a cross-linked gel is obtained by modifying the separation buffer (see Free Solution Capillary Electrophoresis) and by controlling the gel porosity during the gel preparation. For a cross-linked polyacrylamide gel, the porosity can be modified by changing the concentration of acrylamide and/or the ratio of the cross-linker. As a rule, a decrease in the porosity of the gel leads to a decrease in the mobility of the solutes. Due to the rigidity of this type of gel, only electrokinetic injection can be used.

Physical Gels— Physical gels are hydrophilic polymers (i.e., linear polyacrylamide, cellulose derivatives, dextran, etc.) which can be dissolved in aqueous separation buffers, giving rise to a separation medium that also acts as a molecular sieve. These polymeric separation media are easier to prepare than cross-linked polymers. They can be prepared in a vial and filled by pressure in a wall-coated capillary with no electroosmotic flow. Replacing the gel before every injection generally improves the separation reproducibility. The porosity of the physical gels can be increased by using polymers of higher molecular weight (at a given polymer concentration) or by decreasing the polymer concentration (for a given polymer molecular weight). A decrease in gel porosity leads to a decrease in the mobility of the solute for the same buffer. Both hydrodynamic and electromigration injection techniques can be used, since the dissolution of these polymers in the buffer gives low viscosity solutions.

The molecules migrate under the influence of the electric field, so long as they are charged, in a pH gradient generated by ampholytes having pI values in a wide range (poly-aminocarboxylic acids) dissolved in the separation buffer. The three basic steps in capillary isoelectric focusing are loading, focusing, and mobilization.

Loading in One Step— The sample is mixed with ampholytes and introduced into the capillary by pressure or vacuum.

Sequential Loading— A leading buffer, then the ampholytes, then the sample mixed with ampholytes, again ampholytes alone, and finally the terminating buffer are introduced into the capillary. The volume of the sample must be small enough so as to not modify the pH gradient.

Method 1— During Focusing, under the influence of the electroosmotic flow when this flow is small enough to allow the focusing of the components.

Method 2— By application of positive pressure after Focusing.

Method 3— After Focusing, by adding salts in the cathode reservoir or the anode reservoir (depending on the direction chosen for mobilization), in order to alter the pH in the capillary when the voltage is applied. As the pH is changed, the proteins and ampholytes are mobilized in the direction of the reservoir which contains added salts and pass the detector.

Separation takes place in an electrolytic solution which contains a surfactant, generally ionic, at a concentration above the critical micellar concentration. The solute molecules are distributed between the aqueous buffer and the pseudo-stationary phase composed by the micelles according to the solute's partition coefficient. The technique can be considered as a hybrid of electrophoresis and chromatography. It is an electrophoretic technique that can be used for the separation of both neutral and charged solutes maintaining the efficiency, speed, and instrumental suitability of capillary electrophoresis. One of the most widely used surfactants is sodium dodecyl sulfate, although other anionic and cationic surfactants, such as cetyl trimethyl ammonium salts, have also been used.

At neutral and alkaline pH, a strong electroosmotic flow is generated and moves the separation buffer ions in the direction of the cathode. If sodium dodecyl sulfate is used as surfactant, the electrophoretic migration of the anionic micelle is in the opposite direction, toward the anode. As a result, the overall micelle migration velocity is slowed compared to the bulk flow of the electrolytic solution. In the case of neutral solutes, since the analyte can partition between the micelle and the aqueous buffer and has no electrophoretic mobility, the analyte migration velocity will only depend on the partition coefficient between the micelle and the aqueous buffer. In the electrophoretogram, the peak corresponding to each uncharged solute is always between that of the electroosmotic flow marker and that of the micelle, and the time elapsed between these two peaks is called the separation window. For electrically charged solutes, the migration velocity depends on both the partition coefficient of the solute between the micelle and the aqueous buffer and on the electrophoretic mobility of the solute in the absence of micelles.

The separation mechanism is essentially chromatographic, and migration of the solute and resolution can be expressed in terms of the capacity factor of the solute (K¢), which is the ratio between the total number of moles of solute in the micelle to those in the mobile phase. For a neutral compound, K¢ is as follows:

in which tr is the migration time of the solute; to is the analysis time of the unretained solute obtained by injecting an electroosmotic flow marker which does not enter the micelle (i.e., methanol); tm is the micelle migration time measured by injecting a micelle marker, such as Sudan III, which migrates continuously associated in the micelle; K is the partition coefficient of the solute; VS is the volume of the micelles phase; and VM is the volume of the mobile phase.

The resolution between two closely-migrating compounds (RS) is as follows:

in which N is the number of theoretical plates for one of the compounds; is the selectivity obtained; K¢a and K¢b are capacity factors for both components; and the other terms are as defined above.

Similar, but not identical, equations give K¢ and RS values for electrically charged compounds.

INSTRUMENTAL PARAMETERS

Voltage— Separation time is inversely proportional to applied voltage. An increase in voltage can cause excessive heat production that gives rise to temperature gradients and viscosity gradients of the buffer in the cross section of the capillary. This effect can be significant with high conductivity buffers, such as those containing micelles. Poor heat dissipation causes band broadening and decreases resolution.

Temperature— Variations in capillary temperature affect the partition coefficient of the solute between the buffer and the micelle, the critical micelle concentration, and the viscosity of the buffer. These parameters contribute to the migration time of the solutes.

Capillary— Length and internal diameter contribute to analysis time and efficiency of separations. Increasing both effective length and total length can decrease the electrical fields, working at constant voltage, and will increase migration time and improve the separation efficiency. The internal diameter controls heat dissipation, at a given buffer and electrical field, and provides a broadening of the sample band.

ELECTROLYTIC SOLUTION PARAMETERS

Surfactant Type and Concentration— The type of surfactant, as the stationary phase in chromatography, affects the resolution since it modifies separation selectively. The log K¢ of a neutral compound increases linearly with the concentration of detergent in the mobile phase. Resolution in MEKC reaches a maximum when K¢ approaches the value of modifying the concentration of surfactant in the mobile phase changes the resolution.

Buffer pH— pH does not modify the partition coefficient of non-ionized solutes, but it can modify the electroosmotic flow in uncoated capillaries. A decrease in the buffer pH decreases the electroosmotic flow and therefore increases the resolution of the neutral solutes, giving rise to longer analysis time.

Organic Solvents— To improve separation of hydrophobic compounds, organic modifiers (methanol, propanol, acetonitrile, etc.) can be added to the separation electrolytic solution. The addition of these modifiers generally decreases migration time and selectivity of the separation. The addition of organic modifiers affects micelle formation, thus a given surfactant concentration can be used only with a certain percentage of organic modifier before the micellezation equilibrium is eliminated or adversely affected, resulting in the absence of micelles and therefore the absence of the partition mechanism of MEKC. The elimination of micelles in the presence of a high content of organic solvent does not always mean that the separation will no longer be possible, since in some cases, the hydrophobic interaction between the ionic surfactant monomer and the neutral solutes form solvophobic complexes that can be separated electrophoretically.

Additives for Chiral Separations— A chiral selector is included in the micellar system, either covalently bound to the surfactant or added to the micellar separation electrolyte. Micelles which have a moiety with chiral discrimination properties include salts, N-dodecanoyl-L-amino acids, bile salts, etc. Chiral resolution can also be achieved using chiral discriminators, such as cyclodextrins added to the electrolytic solutions which contain micelliced achiral surfactants.

Other Additives— Selectivity can be modified by adding chemicals to the buffer. Addition of several types of cyclodextrins to the buffer are also used to reduce the interaction of hydrophobic solutes with the micelle, increasing the selectivity for this type of compound. The addition of substances able to modify solute-micelle interactions by adsorption on the latter, has been used to improve the selectivity of the separations in MEKC. These additives may consist of a second surfactant (ionic or non-ionic) which gives rise to mixed micelles, metallic cations which dissolve in the micelle, and give co-ordination complexes with the solutes.

QUANTITATIVE ANALYSIS

Calculations— From the values obtained, calculate the content of a component or components being determined. When indicated, the percentage of one (or more) components of the sample to be examined is calculated by determining the areas of the peak(s) as a percentage of the total corrected areas of all the peaks, excluding those due to solvents or any added reagents. The use of an automatic integration system (integrator or data acquisition and processing system) is recommended.

The choice of suitability parameters to be used will depend on the type of capillary electrophoresis that is performed. These parameters are the capacity factor (K¢) used only for Micelles Electrokinetic Chromatography, the number of theoretical plates (n), the symmetry factor (AS), and the resolution (RS). Note that in previous sections, the theoretical expression for n and RS have been described, but more practical equations that allow for the determination of these suitability parameters using the electrophoretograms are described below.

The number of theoretical plates (n) may be calculated from the formula:

in which t is the distance, in mm, along the baseline between the point of injection and the perpendicular dropped from the maximum of the peak in question; and b0.5 is the peak width, in mm, at half height.

The resolution (RS) may be calculated from the formula:

in which tb and ta are the distances, in mm, along the baseline, between the point of injection and the perpendicular dropped from the maxima of two adjacent peaks (tb > ta); and b0.5b and b0.5a are the peak widths, in mm, at half height.

The resolution (RS) may be also calculated by measuring the height of the valley (c) between two partly resolved peaks in a standard preparation, the height of the smaller peak (d), and by specifying (c/d)x, in which x is the limit indicated in the individual monograph.

The symmetry factor of a peak (AS) may be calculated using the formula:

in which b0.05 is the width of the peak at one-twentieth of the peak height; and A is the distance between the perpendicular dropped from the peak maximum and the leading edge of the peak at one-twentieth of the peak height.

Other system suitability parameters include tests for area repeatability (i.e., standard deviation of areas or of area/migration time) and tests for migration time repeatability (i.e., standard deviation of migration time). For migration time repeatability, it will be necessary to provide for a test to measure the suitability of the capillary washing procedures. To avoid the lack of repeatability of the migration time, an alternative practice is to use a migration time relative to an internal standard.

A test for the verification of the signal-to-noise ratio for a standard preparation or the determination of the limit of quantitation is a useful system suitability parameter. The detection limit and quantitation limit correspond to a signal-to-noise ratio greater than 3 and 10, respectively. The signal-to-noise ratio (S/N) is calculated as follows:

in which H is the height of the peak corresponding to the component concerned in the electrophoretogram obtained with the specified reference solution; and hn is the absolute value of the largest noise fluctuation from the baseline in an electrophoretogram obtained after injection of a blank and observed over a distance equal to twenty times the width at the half-height of the peak in the electrophoretogram obtained with the reference solution, and situated equally around the place where this peak would be found.

When a protein is at the position of its isoelectric point, it has no net charge and cannot be moved in a gel matrix by the electric field. It may, however, move from that position by diffusion. The pH gradient forces a protein to remain in its isoelectric point position, thus concentrating it; this concentration effect is called ``focusing''. Increasing the applied voltage or reducing the sample load results in improved resolution of bands. The applied voltage is limited by the heat generated because the heat must be dissipated. The use of thin gels and an efficient cooling plate controlled by a thermostatic circulator prevents the burning of the gel while allowing sharp focusing. The separation is estimated by determining the minimum pI difference, which is necessary to separate two neighboring bands, as follows:

in which D is the diffusion coefficient of the protein; dpH/dx is the pH gradient; E is the intensity of the electric field, in volts per centimeter; and –dµ/dpH is the variation of the solute mobility with the pH in the region close to the pI. Since D and –dµ/dpH for a given protein cannot be altered, the separation can be improved by using a narrower pH range and by increasing the intensity of the electric field.

From an operational point, special attention must be paid to sample characteristics and/or preparation. Salt in a sample can be problematic and it is best to prepare the sample, if possible, in deionized water or 2% ampholytes using dialysis or gel filtration if necessary. Potentials of 2500 volts have been used and are considered optimal under a given set of conditions. Up to 30 watts of constant power can be applied and will generally give complete separation in 1.5 to 3.0 hours. The time required for completion of focusing in thin-layer polyacrylamide gels is determined by placing a colored protein (e.g., hemoglobin) at different positions on the gel surface and by applying the electric field: the steady state is reached when all applications give an identical band pattern. In some procedures the completion of the focusing is indicated by the time elapsed after the sample application.

Resolution between protein bands on an IEF gel prepared with carrier ampholytes can be quite good. Better resolution may be achieved by using immobilized pH gradients where the buffering species, which are analogous to carrier ampholytes, are copolymerized within the gel matrix. Proteins exhibiting pls differing by as little as 0.02 pH units may be resolved using a gel prepared with carrier ampholytes, while immobilized pH gradients can resolve protein differing by approximately 0.001 pH units.

The IEF gel can be used as an identity test when migration on the gel is compared to a standard preparation and IEF calibration proteins, the IEF gel can be used as a limit test when the density of a band on IEF is compared subjectively with the density of bands appearing in a standard preparation, or it can be used as a semi-quantitative test when the density is measured using a densitometer or similar instrumentation to determine the relative concentration of protein in the bands.

An apparatus for isoelectric focusing consists of a controllable direct current generator, of stabilized output; a rigid plastic isoelectric focusing chamber that contains a cooled plate of suitable material to support the gel; and a plastic cover with platinum electrodes that are connected to the gel by means of paper wicks of suitable width, length, and thickness, impregnated with solutions of anodic and cathodic electrolytes.

Unless otherwise indicated in a given monograph, the following procedure in thick polyacrylamide slab gels is to be used.

Assembly— Composed of a glass plate (A) on which a polyester film (B) is placed to facilitate handling of the gel, one or more spacers (C), a second glass plate (D), and clamps to hold the structure together (see Figure 1).

7.5% Polyacrylamide Gel— Dissolve 29.1 g of acrylamide and 0.9 g of methylenebisacrylamide in 100 mL of water. To 2.5 volumes of this solution, add the mixture of ampholytes specified in the individual monograph, and make up to 10 volumes with water. Mix carefully, and degas the solution.

Preparation of the Assembly— Place the polyester film on the lower glass plate, apply the spacer, place the second glass plate, and fit the clamps. Before use, place the mixture on a magnetic stirrer, and add 0.25 volumes of a 10% solution of ammonium persulfate and 0.25 volumes of tetramethylenediamine. Immediately fill the space between the glass plates of the assembly with the gel.

Fixing Solution for Isoelectric Focusing Polyacrylamide Gel— Mix 35 g of sulfosalicylic acid and 100 g of trichloroacetic acid in 1000 mL of water.

Coomassie Staining Solution and Destaining Solution— Use the same solutions indicated in Polyacrylamide Gel Electrophoresis.

Procedure— Dismantle the assembly, and using the polyester film, transfer the gel onto the cooled support wetted with a few milliliters of a suitable liquid, taking care to avoid forming air bubbles. Prepare the test solutions and reference solutions as specified in the individual monograph. Place strips of paper for sample application, about 10 mm × 5 mm in size, on the gel, and impregnate each with the prescribed amount of the test and reference solutions. If the protein concentration of the solution is too low, several strips may be superimposed (up to four). Also apply the prescribed quantity of a solution of proteins with known isoelectric points as pH markers to calibrate the gel. In some procedures, the gel has pre-cast slots where a solution of the sample is applied instead of using impregnated paper strips. Cut two strips of paper to the length of the gel, and impregnate them with the electrolyte solutions: acid for the anode and alkaline for the cathode. The compositions of the anode and cathode solutions are given in the individual monograph. Apply these paper wicks to each side of the gel several millimeters from the edge. Fit the cover so that the electrodes are in contact with the wicks (with respect to the anodic and cathodic poles). Proceed with the isoelectric focusing by applying the electrical parameters described in the individual monograph. Switch off the current when the migration of the mixture of standard proteins has stabilized. Using forceps, remove the sample application strips and the two electrode wicks. Immerse the gel in Fixing Solution for Isoelectric Focusing Polyacrylamide Gel. Incubate with gentle shaking at room temperature for 30 minutes. Drain off the solution, and add 200 mL of Destaining Solution. Incubate with shaking for 1 hour. Drain the gel, add Coomassie Staining Solution. Incubate for 30 minutes. Destain the gel by passive diffusion with Destaining Solution until the bands are well visualized against a clear background. Locate the position and intensity of the bands in the electropherogram as prescribed in the individual monograph.

Alternative Procedure— When a monograph references the general method for isoelectric focusing above, variations in methodology or procedure may be used, subject to validation. These variations include the use of commercially available pre-cast gels; the use of immobilized pH gradients; the use of rod gels; and the use of cassettes of different dimensions, including ultra-thin (0.2 mm) gels; the variations in the sample application procedure, including different sample volumes or the use of sample application masks or wicks other than paper; the use of alternate running conditions, including variations in the electric field depending on gel dimensions and equipment, and the use of fixed migration times rather than subjective interpretation of band stability; the inclusion of a pre-focusing step; the use of automated instrumentation; and the use of agarose gels.

Where alternative methods to the general method are employed, they must be validated. The following criteria may be used to validate the separation: the formation of a stable pH gradient of desired characteristics, evaluated using colored pH markers of known isoelectric points; the comparison with electropherogram provided with the chemical reference substance for the preparation to be examined; and any other validation criteria as prescribed in the individual monograph.

Variations to the general method required for the analysis of specific substances may be specified in detail in individual monographs. Variations may include the addition of urea in the running gel (3 M concentration is often satisfactory to keep protein in solution but up to 8 M can be used). Some proteins precipitate at their isoelectric point. In this case, urea is included in the gel formulation to keep the protein in solution. If urea is used, only fresh solutions should be used to prevent carbamylation of the protein. Other variations include the use of alternative staining methods and the use of gel additives such as non-ionic detergents (e.g., octylglucoside) or zwitterionic detergents (e.g., CHAPS or CHAPSO) to prevent proteins from aggregating or precipitating.

Peptide mapping is an identity test for proteins, especially those obtained by r-DNA technology. It involves the chemical or enzymatic treatment of a protein resulting in the formation of peptide fragments followed by separation and identification of the resultant fragments in a reproducible manner. It is a powerful test that is capable of identifying single amino acid changes resulting from events such as errors in the reading of complementary DNA (cDNA) sequences or point mutations. Peptide mapping is a comparative procedure because the information obtained, compared to a reference standard or reference material similarly treated, confirms the primary structure of the protein, is capable of detecting whether alterations in structure have occurred, and demonstrates process consistency and genetic stability. Each protein presents unique characteristics which must be well understood so that the scientific and analytical approaches permit validated development of a peptide map that provides sufficient specificity.

This section provides detailed assistance in the application of peptide mapping and its validation to characterize the desired protein product, to evaluate the stability of the expression construct of cells used for recombinant DNA products, to evaluate the consistency of the overall process, and to assess product stability, as well as to ensure the identity of the protein product, or to detect the presence of protein variant. The validation scheme presented differentiates between qualification of the method at an early stage in the regulatory process, Investigational New Drug (IND) level, and full validation in support of New Drug Application (NDA), Product License Application (PLA), or Marketing Authorization Application (MAA). The validation concepts described are consistent with the general information chapter Validation of Compendial Methods 1225 and with the International Conference on Harmonization (ICH) document on Analytical Methods Validation.

Peptide mapping is not a general method, but involves developing specific maps for each unique protein. Although the technology is evolving rapidly, there are certain methods that are generally accepted. Variations of these methods will be indicated, when appropriate, in specific monographs.

A peptide map may be viewed as a fingerprint of a protein and is the end product of several chemical processes that provide a comprehensive understanding of the protein being analyzed. Four major steps are necessary for the development of the procedure: isolation and purification of the protein, if the protein is part of a formulation; selective cleavage of the peptide bands; chromatographic separation of the peptides; and analysis and identification of the peptides. A test sample is digested and assayed in parallel with a reference standard or reference material. Complete cleavage is more likely to occur when enzymes such as endoproteases (e.g., trypsin) are used instead of chemical cleavage reagents. A map should contain enough peptides to be meaningful. On the other hand, if there are too many fragments, the map might lose its specificity because many proteins will then have the same profiles.

ISOLATION and PURIFICATION

SELECTIVE CLEAVAGE OF PEPTIDE BONDS

Type	Agent	Specificity
Enzymatic	Trypsin (EC 3.4.21.4)	C-terminal side of Arg and Lys
	Chymrotrypsin (EC 3.4.21.1)	C-terminal side of hydrophobic residues (e.g., Leu, Met, Ala, aromatics)
	Pepsin (EC 3.4.23.122)	Nonspecific digest
	Lysyl endopeptidase (Lys-C Endopeptidase)	C-terminal side of Lys
	(EC 3.4.21.50)
	Glutamyl endopeptidase (from S. aureus stain V8)	C-terminal side of Glu and Asp
	(EC 23.4.21.19)
	Peptidyl-Asp metaplo-endopeptidase	N-terminal side of Asp
	(Endoproteinase Asp-N)
	(EC 3.4.24.33)
	(Clostripan)	C-terminal side of Arg
	(EC 3.4.28.8)
Chemical	Cyanogen bromide	C-terminal side of Met
	2-Nitro-5-thio-cyano- benzoic acid	N-terminal side of Cys
	O-Iodosobenzoic acid	C-terminal side of Trp and Tyr
	Dilute acid	Asp and Pro
	BNPS-skatole	Trp

Pretreatment of Sample— Depending on the size or the configuration of the protein, different approaches in the pretreatment of samples can be used. For monoclonal antibodies, the heavy and light chains will need to be separated before mapping. If trypsin is used as a cleavage agent for proteins with a molecular mass greater than 100,000 Da, lysine residues must be protected by citraconylation or maleylation; otherwise, too many peptides will be generated.

Pretreatment of the Cleavage Agent— Pretreatment of cleavage agents—especially enzymatic agents—might be necessary for purification purposes to ensure reproducibility of the map. For example, trypsin used as a cleavage agent will have to be treated with tosyl-L-phenylalanine chloromethyl ketone to inactivate chymotrypsin. Other methods, such as purification of trypsin by HPLC or immobilization of enzyme on a gel support, have been successfully used when only a small amount of protein is available.

Pretreatment of the Protein— Under certain conditions, it might be necessary to concentrate the sample or to separate the protein from added substances and stabilizers used in the formulation of the product, if these interfere with the mapping procedure. Physical procedures used for pretreatment can include ultrafiltration, column chromatography, and lyophilization.

Establishment of Optimal Digestion Conditions— Factors that affect the completeness and effectiveness of digestion of proteins are those that could affect any chemical or enzymatic reactions.

CHROMATOGRAPHIC SEPARATION

Reverse-Phase High-Performance Liquid

Chromatography (RP-HPLC)

Ion-Exchange Chromatography (IEC)

Hydrophobic Interaction Chromatography (HIC)

Polyacrylamide Gel Electrophoresis (PAGE), nondenaturating

Sodium Dodecyl Sulfate Polyacrylamide Gel

Electrophoresis (SDS-PAGE)

Capillary Electrophoresis (CE)

Paper Chromatography

High-Voltage Paper Electrophoresis (HVPE)

Chromatographic Column— The selection of a chromatographic column is empirically determined for each protein. Columns with 100 or 300 pore size with silica support can give optimal separation. For smaller peptides, octylsilane chemically bonded to totally porous silica articles, 3 to 10 µm in diameter (L7) and octadecylsilane chemically bonded to porous silica or ceramic microparticles, 3 to 10 µm in diameter (L1) column packings are more efficient than the butyl silane chemically bonded to totally porous silica particles, 5 to 10 µm in diameter (L26) packing.

Solvent— The most commonly used solvent is water with acetonitrile as the organic modifier to which less than 0.1% of trifluoroacetic acid is added. If necessary, add isopropyl alcohol or n-propyl alcohol to solubilize the digest components, provided that the addition does not unduly increase the viscosity of the components.

Mobile Phase— Buffered mobile phases containing phosphate are used to provide some flexibility in the selection of pH conditions, since shifts of pH in the 3.0 to 5.0 range enhance the separation of peptides containing acidic residues (e.g., glutamic and aspartic acids). Sodium or potassium phosphates, ammonium acetate, phosphoric acid, and a pH between 2 and 7 (or higher for polymer-based supports) have also been used with acetonitrile gradients. Acetonitrile-containing trifluoroacetic acid is also used quite often.

Gradient Selection— Gradients can be linear, nonlinear, or include step functions. A shallow gradient is recommended in order to separate complex mixtures. Gradients are optimized to provide clear resolution of one or two peaks that will become “marker” peaks for the test.

Isocratic Selection— Isocratic HPLC systems using a single mobile phase are used on the basis of their convenience of use and improved detector responses. Optimal composition of a mobile phase to obtain clear resolution of each peak is sometimes difficult to establish. Mobile phases for which slight changes in component ratios or in pH significantly affect retention times of peaks in peptide maps should not be used in isocratic HPLC systems.

Other Parameters— Temperature control of the column is usually necessary to achieve good reproducibility. The flow rates for the mobile phases range from 0.1 to 2.0 mL per minute, and the detection of peptides is performed with a UV detector at 200 to 230 nm. Other methods of detection have been used (e.g., postcolumn derivatization), but they are not as robust or as versatile as UV detection.

System Suitability— The section System Suitability under Chromatography 621 provides an experimental means for measuring the overall performance of the test method. The acceptance criteria for system suitability depend on the identification of critical test parameters that affect data interpretation and acceptance. These critical parameters are also criteria that monitor peptide digestion and peptide analysis. An indicator that the desired digestion endpoint was achieved is the comparison with a reference standard or reference material, which is treated exactly as the article under test. The use of a USP Reference Standard in parallel with the protein under test is critical in the development and establishment of system suitability limits. In addition, a specimen chromatogram should be included with the USP Reference Standard or reference material for comparison purposes. Other indicators may include visual inspection of protein or peptide solubility, the absence of intact protein, or measurement of responses of a digestion-dependent peptide. The critical system suitability parameters for peptide analysis will depend on the particular mode of peptide separation and detection, and on the data analysis requirements.

ANALYSIS and IDENTIFICATION OF PEPTIDES

A validated peptide map can be used to assess the integrity of the predicted primary sequence of a protein product (i.e., its genetic stability). It can also be used to determine lot-to-lot consistency of the biotechnology-derived product process. Furthermore, the performance of the protein expression of the production system is best assessed by peptide mapping of the expressed protein. Peptide maps of protein produced at various times of the protein expression process, including a point well beyond the normal protein expression time, compared with those of a USP Reference Standard or reference material, will evaluate the genetic stability of the expression system as a function of time.

Variant protein sequences can arise from a genetic variation at the DNA level (point mutation) or as an error in the translation process. A validated peptide map is the best approach to the detection of protein variants. However, the limitations of the peptide mapping itself must be taken into consideration. The detection of a structured variant is possible only if the corresponding peptide variant is easily isolated and characterized. To establish genetic stability will require the use of a battery of biochemical methods, provided that the variants have properties different from those of the “normal” protein.

CRITICAL FACTORS

Written Test Procedures— These procedures include a detailed description of the analytical method in which reagents, equipment, sample preparation, method of analysis, and analysis of the data are defined.

Validation Protocol— A protocol is prepared that contains a procedure for test validation.

Acceptance Criteria— The criteria can be minimal at the early stages, but need to be better defined as validation studies progress.

Reporting of Results— Results from the validation study are documented with respect to the analytical parameters listed in the validation protocol.

Revalidation of the Test Procedure— If the method used requires alteration that could affect the analytical parameter previously assessed in the validation of the procedure, the test procedure must be revalidated. Significant changes in the processing of the article, in laboratories performing the analysis, in formulation of the bulk or the finished products, and in any other significant parameter will require revalidation of the methods.

REQUIREMENTS

Precision—

Robustness— Factors such as composition of the Mobile Phase, protease quality or chemical reagent purity, column variation and age, and digest stability are likely to affect the overall performance of the test and its reproducibility. Tolerances for each of the key parameters are evaluated and baseline limits established in case the test is used for routine lot release purposes.

Reproducibility— Determination of various parameters indicated above is repeated using the same USP Reference Standard or reference material and test sample in at least two different laboratories by two analysts equipped with similar HPLC systems. The generated peptide maps are evaluated for significant differences.

Under the influence of an electrical field, charged particles migrate in the direction of the electrode bearing the opposite polarity. In gel electrophoresis, the movements of the particles are retarded by interactions with the surrounding gel matrix, which acts as a molecular sieve. The opposing interactions of the electrical force and molecular sieving result in differential migration rates according to sizes, shapes, and charges of particles. Because of their different physicochemical properties, different macromolecules of a mixture will migrate at different speeds during electrophoresis and thus will be separated into discrete fractions. Electrophoretic separations can be conducted in systems without support phases (e.g., free solution separation in capillary electrophoresis) and in stabilizing media, such as thin-layer plates, films, or gels.

The sieving properties of polyacrylamide gels are established by the three-dimensional network of fibers and pores that is formed as the bifunctional bisacrylamide cross-links adjacent to polyacrylamide chains. Polymerization is catalyzed by a free radical-generating system composed of ammonium persulfate and N,N,N¢,N¢-tetramethylethylenediamine (TEMED).

As the acrylamide concentration of a gel increases, its effective pore size decreases. The effective pore size of a gel is operationally defined by its sieving properties, that is, by the resistance it imparts to the migration of macromolecules. There are limits to the acrylamide concentrations that can be used. At high acrylamide concentrations, gels break much more easily and are difficult to handle. As the pore size of a gel decreases, the migration rate of a protein through the gel decreases. By adjusting the pore size of a gel, through manipulating the acrylamide concentration, the resolution of the method can be optimized for a given protein product. Thus, a given gel is physically characterized by its respective composition of acrylamide and bisacrylamide.

In addition to the composition of the gel, the state of the protein is an important component to the electrophoretic mobility. In the case of proteins, the electrophoretic mobility is dependent on the pK value of the charged groups and the size of the molecule. It is influenced by the type, the concentration, and pH of the buffer, by the temperature and the field strength, and by the nature of the support material.

Denaturing PAGE using sodium dodecyl sulfate (SDS) is the most common mode of electrophoresis used in assessing the pharmaceutical quality of protein products. Typically, analytical electrophoresis of proteins is carried out under conditions that ensure dissociation of the proteins into their individual polypeptide subunits and that minimize aggregation of these subunits. The strongly anionic detergent SDS is used in combination with heat to dissociate the proteins before they are loaded on the gel. The denatured polypeptides bind SDS, become negatively charged, and exhibit a consistent charge-to-weight ratio regardless of protein type. Because the amount of SDS bound is almost always proportional to the molecular weight of the polypeptide and is typically independent of its sequence, SDS-polypeptide complexes migrate through polyacrylamide gels in reasonable accordance with the size of the polypeptide.

The electrophoretic mobilities of the resultant detergent-polypeptide complexes all assume the same functional relationship to the molecular weights of polypeptides. Migration of SDS derivatives is toward the anode in a predictable manner, with low molecular weight complexes migrating faster than larger ones. This means that the molecular weight of a protein can be estimated from its relative mobility in calibrated SDS-PAGE and that a single band in such a gel is a criterion of purity.

Modifications to the polypeptide backbone, such as N- or O-linked glycosylation, however, have a significant impact on the apparent molecular weight of a protein. This is due to SDS not binding to a carbohydrate moiety in a manner similar to the polypeptide. Thus, a consistent charge-to-weight ratio is not maintained. The apparent molecular weight of proteins having undergone post-translational modifications is not a true reflection of the weight of the polypeptide chain.

REDUCING CONDITIONS

NONREDUCING CONDITIONS

CHARACTERISTICS OF A DISCONTINUOUS BUFFER SYSTEM

PREPARATION OF GELS

Gel Stock Solutions—

Plate Preparation— Clean two glass plates (10 cm × 8 cm), the polytef comb, the two spacers, and the silicone rubber tubing (0.6 mm × 35 cm) with mild detergent, rinse thoroughly with water, and blot dry.

Resolving Gel— In a conical flask, prepare the appropriate volume of solution containing the desired concentration of acrylamide as directed in Table 3. Mix the components in the order shown. Before adding the Ammonium Persulfate Solution and the TEMED, pour the solution in a disposable filtration unit equipped with a nitrocellulose filter having a 0.45-µm porosity, and apply vacuum. Allow the solution to degas by swirling the filtration unit, and disconnect the vacuum when no more bubbles are formed in the solution. Add appropriate amounts of Ammonium Persulfate Solution and TEMED, as directed in Table 3, swirl, and pour immediately into the gap between the two glass plates of the mold. Leave sufficient space for the stacking gel (the length of the teeth of the comb plus 1 cm). Using a pipet, carefully overlay the solution with water-saturated isobutyl alcohol. Leave the gel in a vertical position at room temperature for polymerization.

Stacking Gel— In a conical flask, prepare the appropriate volume of solution containing the desired concentration of acrylamide, as directed in Table 4. Mix the components in the order shown. Before adding the Ammonium Persulfate Solution and the TEMED, pour the solution into a disposable filtration unit equipped with a nitrocellulose filter having a 0.45-µm porosity, and apply vacuum. Allow the solution to degas by swirling the filtration unit, and disconnect the vacuum when no more bubbles are formed in the solution. Add appropriate amounts of Ammonium Persulfate Solution and TEMED as directed in Table 4, swirl, and pour immediately into the gap between the two glass plates of the mold directly onto the surface of the polymerized Resolving Gel. Immediately insert a clean polytef comb into the stacking gel solution, being careful to avoid trapping air bubbles. Add more stacking gel solution to fill the spaces of the comb completely. Leave the gel in a vertical position, and allow to polymerize at room temperature. After polymerization is complete (about 30 minutes later), carefully remove the polytef comb, and proceed as directed below.

ELECTROPHORETIC SEPARATION

Sample Buffer 1— Dissolve 1.89 g of TRIS, 5.0 g of SDS, 50 mg bromophenol blue, and 25.0 mL glycerol in 100 mL of water. Adjust with hydrochloric acid to a pH of 6.8, and dilute with water to 125 mL. Before use, dilute with an equal volume of water or sample, and mix.

Sample Buffer 2 (for reducing conditions)— Prepare as directed under Sample Buffer 1 except to add 12.5 mL of 2-mercaptoethanol before adjusting the pH. Alternatively, prepare as directed for Sample Buffer 1 except to start with about 1.93 g of TRIS and add a suitable quantity of DTT to obtain a final 100 µM DTT concentration.

Running Buffer— Dissolve 151.4 g of TRIS, 721.0 g of aminoacetic acid, and 50.0 g of SDS in water, dilute with water to 5000 mL, and mix to obtain a stock solution. Immediately before use, dilute this stock solution with water to 10 times its volume, mix, and adjust to a pH between 8.1 and 8.8.

Procedure— Rinse the wells immediately with water or with the Running Buffer to remove any unpolymerized acrylamide. (If necessary, straighten the teeth of the Stacking Gel with a blunt hypodermic needle attached to a syringe.) Remove the clamps on one short side, carefully pull out the tubing, and replace the clamps. Proceed similarly on the other short side. Remove the tubing from the bottom part of the gel.

DETECTION OF PROTEINS IN GELS

Reagents—

Coomassie Staining— Immerse the gel in an excess of Coomassie Staining Solution, and incubate for at least 1 hour. Remove the Coomassie Staining Solution. Destain the gel with an excess of Destaining Solution. Change the Destaining Solution several times, until the stained protein bands are clearly distinguishable on a clear background. The more thoroughly the gel is destained, the smaller the amount of protein that can be detected. Destaining can be accelerated by including a few g of anion-exchange resin or a small sponge in the Destaining Solution. [NOTE—The acid-alcohol solutions used in this procedure do not completely fix proteins in the gel. This can lead to losses of some low molecular weight proteins during the staining and destaining of thin gels. Permanent fixation is obtainable by incubating the gel in Fixing Solution 1 for 1 hour before it is immersed in the Coomassie Staining Solution.]

Silver Staining— Immerse the gel in an excess of Fixing Solution 2, and incubate for 1 hour. Remove Fixing Solution 2, add fresh Fixing Solution 2, and incubate for at least 1 hour, or overnight if convenient. Discard Fixing Solution 2, and wash the gel in an excess of water for 1 hour. Soak the gel for 15 minutes in a 1% solution of glutaraldehyde (v/v). Wash the gel twice, for 15 minutes each time, with an excess of water. Soak the gel in fresh Silver Nitrate Reagent for 15 minutes, in darkness. Wash the gel three times, for 5 minutes each time, with an excess of water. Immerse the gel for about 1 minute in Developing Solution until satisfactory staining has been obtained. Stop the development by incubation in the Stopping Solution for 15 minutes, then rinse the gel with water, and proceed with drying as indicated below.

DRYING OF GELS

MOLECULAR WEIGHT DETERMINATION

QUANTIFICATION OF IMPURITIES

Protein in solution absorbs UV light at a wavelength of 280 nm, due to the presence of aromatic amino acids, mainly tyrosine and tryptophan. This property is the basis of this method. Protein determination at 280 nm is mainly a function of the tyrosine and tryptophan content of the protein. If the buffer used to dissolve the protein has a high absorbance relative to that of water, there is an interfering substance in the buffer. This interference can be compensated for when the spectrophotometer is adjusted to zero buffer absorbance. If the interference results in a large absorbance that challenges the limit of sensitivity of the spectrophotometer, the results may be compromised. Furthermore, at low concentrations protein can be absorbed onto the cuvette, thereby reducing the content in solution. This can be prevented by preparing samples at higher concentrations or by using a nonionic detergent in the preparation. [NOTE—Keep the Test Solution, the Standard Solution, and the buffer at the same temperature during testing.]

CS(AU / AS),

This method, commonly referred to as the Lowry assay, is based on the reduction by protein of the phosphomolybdic-tungstic mixed acid chromogen in the Folin-Ciocalteu's phenol reagent, resulting in an absorbance maximum at 750 nm. The Folin-Ciocalteu's phenol reagent reacts primarily with tyrosine residues in the protein, which can lead to variation in the response of the assay to different proteins. Because the method is sensitive to interfering substances, a procedure for precipitation of the protein from the test specimen may be used. Where separation of interfering substances from the protein in the test specimen is necessary, proceed as directed below for Interfering Substances prior to preparation of the Test Solution. The effect of interfering substances can be minimized by dilution, provided the concentration of the protein under test remains sufficient for accurate measurement.

Copper Sulfate Reagent— Dissolve 100 mg of cupric sulfate and 200 mg of sodium tartrate in water, dilute with water to 50 mL, and mix. Dissolve 10 g of sodium carbonate in water to a final volume of 50 mL, and mix. Slowly pour the sodium carbonate solution into the copper sulfate solution with mixing. Prepare this solution fresh daily.

SDS Solution— Dissolve 5 g of sodium dodecyl sulfate in water, and dilute with water to 100 mL.

Sodium Hydroxide Solution— Dissolve 3.2 g of sodium hydroxide in water, dilute with water to 100 mL, and mix.

Alkaline Copper Reagent— Prepare a mixture of Copper Sulfate Reagent, SDS Solution, and Sodium Hydroxide Solution (1:2:1). This reagent may be stored at room temperature for up to 2 weeks.

Diluted Folin-Ciocalteu’s Phenol Reagent— Mix 10 mL of Folin-Ciocalteu’s phenol TS with 50 mL of water. Store in an amber bottle, at room temperature.

INTERFERING SUBSTANCES

Sodium Deoxycholate Reagent— Prepare a solution of sodium deoxycholate in water having a concentration of 150 mg in 100 mL.

Trichloroacetic Acid Reagent— Prepare a solution of trichloroacetic acid in water having a concentration of 72 g in 100 mL.

Procedure— Add 0.1 mL of Sodium Deoxycholate Reagent to 1 mL of a solution of the protein under test. Mix on a vortex mixer, and allow to stand at room temperature for 10 minutes. Add 0.1 mL of Trichloroacetic Acid Reagent, and mix on a vortex mixer. Centrifuge at 3000 × g for 30 minutes, decant the liquid, and remove any residual liquid with a pipet. Redissolve the protein pellet in 1 mL of Alkaline Copper Reagent. Proceed as directed for the Test Solution.

This method, commonly referred to as the Bradford assay, is based on the absorption shift from 470 nm to 595 nm observed when the brilliant blue G dye binds to protein. The brilliant blue G dye binds most readily to arginyl and lysyl residues in the protein, which can lead to variation in the response of the assay to different proteins.

This method, commonly referred to as the bicinchoninic acid or BCA assay, is based on reduction of the cupric (Cu2+) ion to cuprous (Cu1+) ion by protein. The bicinchoninic acid reagent is used to detect the cuprous ion. The method has few interfering substances. When interfering substances are present, their effect may be minimized by dilution, provided that the concentration of the protein under test remains sufficient for accurate measurement.

BCA Reagent— Dissolve about 10 g of bicinchoninic acid, 20 g of sodium carbonate monohydrate, 1.6 g of sodium tartrate, 4 g of sodium hydroxide, and 9.5 g of sodium bicarbonate in water. Adjust, if necessary, with sodium hydroxide or sodium bicarbonate to a pH of 11.25. Dilute with water to 1 L, and mix.

Copper Sulfate Reagent— Dissolve about 2 g of cupric sulfate in water to a final volume of 50 mL.

Copper-BCA Reagent— Mix 1 mL of Copper Sulfate Reagent and 50 mL of BCA Reagent.

This method, commonly referred to as the Biuret assay, is based on the interaction of cupric (Cu2+) ion with protein in an alkaline solution and the resultant development of absorbance at 545 nm.

This fluorometric method is based on the derivatization of the protein with o-phthalaldehyde (OPA), which reacts with the primary amines of the protein (i.e., NH2-terminal amino acid and the -amino group of the lysine residues). The sensitivity of the test can be increased by hydrolyzing the protein before testing. Hydrolysis makes the -amino group of the constituent amino acids of the protein available for reaction with the o-phthalaldehyde reagent. The method requires very small quantities of the protein.

Primary amines, such as tris(hydroxymethyl)aminomethane and amino acid buffers, react with o-phthalaldehyde and must be avoided or removed. Ammonia at high concentrations will react with o-phthalaldehyde as well. The fluorescence obtained when amine reacts with o-phthalaldehyde can be unstable. The use of automated procedures to standardize this procedure may improve the accuracy and precision of the test.

Borate Buffer— Dissolve about 61.83 g of boric acid in water, and adjust with potassium hydroxide to a pH of 10.4. Dilute with water to 1 L, and mix.

Stock OPA Reagent— Dissolve about 120 mg of o-phthalaldehyde in 1.5 mL of methanol, add 100 mL of Borate Buffer, and mix. Add 0.6 mL of polyoxyethylene (23) lauryl ether, and mix. This solution is stable at room temperature for at least 3 weeks.

OPA Reagent— To 5 mL of Stock OPA Reagent add 15 µL of 2-mercaptoethanol. Prepare at least 30 minutes prior to use. This reagent is stable for one day.

This method is based on nitrogen analysis as a means of protein determination. Interference caused by the presence of other nitrogen-containing substances in the test specimen can affect the determination of protein by this method. Nitrogen analysis techniques destroy the protein under test but are not limited to protein presentation in an aqueous environment.