The migration of charged colloidal particles or molecules through a solution under the influence of an applied electric field is usually provided by immersed electrodes. It can also be described as a method of separating substances, especially proteins and analyzing molecular structure based on the rate of movement of each component in a colloidal suspension under the influence of an electric field. An analyte is a chemical substance that is the subject of chemical analysis. Separation by electrophoresis depends on differences in the migration velocity of ions or solutes through a given medium in an electric field. The electrophoretic migration velocity of an analyte is where E is the electric field strength and  is the electrophoretic mobility.

The electrophoretic mobility is inversely proportional to frictional forces in the buffer, and directly proportional to the ionic charge of the analyte. The forces of friction against an analyte are dependent on the analyte’s size and the viscosity (η) of the medium. Analytes with different frictional forces or different charges will separate from one another when they move through a buffer. At a given pH, the electrophoretic mobility of an analyte is:
where, r is the radius of the analyte and z is the net charge of the analyte.

Differences in the charge to size ratio of analyte cause differences in electrophoretic mobility. Small, highly charged analytes have greater mobility, whereas large, less charged analytes have lower mobility. Electrophoretic mobility is an indication of an analyte’s migration velocity in a given medium. The net force acting on an analyte is the balance of two forces: the electrical force acting in favor of motion, and the frictional force acting against motion. The two forces remain steady during electrophoresis, thus the electrophoretic mobility is a constant for a given analyte under a given set of conditions.

Electrophoresis has a wide variety of applications in proteomics, forensics, molecular biology, genetics, biochemistry, and microbiology. One of the most common uses of electrophoresis is to analyze differential expression of genes. Healthy and diseased cells can be identified by differences in the electrophoretic patterns of their proteins. Proteins can also be characterized similarly, and some information about their structure can be derived from the masses of fragments in the gel.

Two-dimensional polyacrylamide gel electrophoresis (2DE) was first described by O’Farrell in 1975 and has evolved markedly as one of the core technologies for the analysis of complex protein mixtures extracted from biological samples since then. The proteins are separated in 2 steps according to 2 independent properties [isoelectric point (pI) and molecular weight (MW)].

The proteins are made up of amino acids which may have positive, negative, or no charges. In addition, amino acids can be hydrophobic or hydrophilic. These amino acid properties are combined in the molecule to determine the electrostatic and amphiphilic properties of the proteins. When a protein of charge q is placed in an electric field, E, it experiences an electrical force given by: F = q × E. Under the influence of this force, the protein moves until the force becomes zero (F = 0). This principle is used to make proteins move in a liquid or solid media. A common medium used is Polyacrylamide (PAA). PAA is a flexible, elastic polymer that allows particles to move inversely proportional to their sizes; i.e., smaller particles move faster than larger particles. Thus a mixture of proteins in a PAA gel under the influence of a force F will separate in individual molecules depending on their sizes and charges. Two-dimensional polyacrylamide gel electrophoresis (2D-PAGE) is a form of gel electrophoresis in which proteins are separated and identified in two dimensions oriented at right angles to each other. Small changes in charge and mass can easily be detected by this method, because it is rare that two different proteins will resolve to the same place in both dimensions. A 2D gel can resolve one thousand to two thousand proteins, which appear, after staining, as dots in the gel.

The three main advantages of this technique are its robustness, its parallelism, and its unique ability to analyze complete proteins at high resolution. This technique is useful when comparing two similar samples to find specific protein differences. Using 2D-PAGE, hundreds to thousands of polypeptides can be analyzed in a single run. The proteins can be separated in pure form from the resultant spots which can be quantified and further analyzed by mass spectrometry, depending on their resolution. Polypeptides can also be probed with antibodies and tested for post-translational modifications. 2D-PAGE is also used to study differential expression of proteins between cell types. The two main drawbacks are its very low efficiency in the analysis of hydrophobic proteins, and high sensitivity toward the dynamic range and quantitative distribution issues. It requires a large amount of sample handling, limited reproducibility, and a smaller dynamic range than other separation methods. It is also not automated for high-throughput analysis. Certain proteins are difficult for 2D-PAGE to separate such as those that are in low abundance, acidic, basic, hydrophobic, very large, or very small.

Multi-dimentional protein identification technology (MudPIT) is a chromatography-based proteomic technique in which a complex peptide mixture is prepared from a protein sample and loaded directly onto a triphasic microcapillary column packed with reversed phase, strong cation exchange, and reversed phase HPLC grade materials. Once the complex peptide mixture is loaded the column is placed directly in-line with a tandem mass spectrometry (MS/MS). The MS/MS data generated from a MudPIT run is then searched to determine the protein content of the original sample.

MudPIT is a robust and widely accepted method for protein identification from a wide variety of samples. It is an excellent tool for both qualitative and quantitative proteomic analyses. Through on-line 2D HPLC, complex peptides mixtures can be well separated. For a relatively simple sample, better results are obtained from MudPIT, compared to single dimension liquid chromatography method. It was developed as a method to analyze the highly complex samples necessary for large-scale proteome analysis by electrospray ionization, MS/MS, and database searching. This method couples a 2D liquid chromatography separation of peptides on a microcapillary column with detection in a tandem mass spectrometer. In the MudPIT, a protein or mixture of proteins is first reduced (to break cysteine disulfide bonds), alkylated (to prevent reformation of disulfide bonds), and digested into a complex mixture of peptides.

MudPIT has been used in a wide range of proteomics experiments, including large-scale catalogs of proteins in cells and organisms, profiling of organelle and membrane proteins, identification of protein complexes, determination of post-translational modifications and quantitative analysis of protein expression. In MudPIT biochemical fractions containing many proteins are directly proteolyzed and the enormous number of peptides generated, are separated by 2D liquid chromatography before entering the mass spectrometer. Instead of MALDI-TOF, MS/MS is employed so that, after the mass of a peptide is measured, the peptide is fragmented using a collision-induced dissociation cell, and the masses of the fragmentation products are determined.

Isotope-coded affinity tags (ICATs) are gel-free method for quantitative proteomics that relies on chemical labeling reagents. These chemical probes consist of three general elements: a reactive group capable of labeling a defined amino acid side chain (e.g., iodoacetamide to modify cysteine residues), an isotopically coded linker and a tag (e.g., biotin) for the affinity isolation of labeled proteins/peptides. For the quantitative comparison of two proteomes, one sample is labeled with the isotopically light (d0) probe and the other with the isotopically heavy (d8) version (Fig. 8.5). To minimize error, both samples are then combined, digested with a protease (i.e., trypsin), and subjected to avidin affinity chromatography to isolate peptides labeled with isotope-coded tagging reagents. These peptides are then analyzed by liquid chromatography–mass spectrometry (LC–MS). The ratios of signal intensities of differentially mass-tagged peptide pairs are quantified to determine the relative levels of proteins in the two samples. The original tags were developed using deuterium, but later tags using ¹³C were used instead to circumvent issues of peak separation during liquid chromatography due to the interaction of deuterium with the stationary phase of the column.

For samples that are not amenable to metabolic labeling, such as when analyzing clinical samples (e.g., biological fluids, tissue samples) or when experimental time is limited, chemical or enzymatic stable isotopic labeling methods are available for quantitative proteomic analyses. These include strategies to add isotopic atoms or isotope-coded tags to peptides or proteins. A rapid and relatively inexpensive method of chemical labeling is stable isotope dimethylation which uses formaldehyde in deuterated water to label primary amines with deuterated methyl groups. This approach also does not change the ionic state of the labeled peptides because of the reductive amination that occurs, so their chemical properties remain the same as those of unlabeled peptides. Benefit of this approach is that many samples are amenable to formaldehyde fixation, which is fast and cheap compared to other labeling reagents. This requires using pure samples or sample preparation to reduce the complexity of biological samples to minimize the number of peaks detected by MS.

Protein labeling with ICAT followed by MS/MS allows sequence identification and accurate quantification of proteins in complex mixtures, and has been applied to the analysis of global protein expression changes, protein changes in subcellular fractions, components of protein complexes, protein secretion, and body fluids.

Label-free methods for both relative and absolute quantitation have been developed as a rapid and low-cost alternative to other quantitative proteomic approaches. These strategies are ideal for large-sample analyses in clinical screening or biomarker discovery experiments but are less reliable for measuring small changes. Unlike other quantitation methods, label-free samples are separately collected, prepared, and analyzed by liquid chromatography–mass spectrometry, LC–MS, or LC–MS/MS. Hence, label-free quantitation experiments need to be more carefully controlled than stable isotope methods to account for any experimental variations. Protein quantitation is performed using either ion peak intensity or spectral counting.

Relative quantitation by ion peak intensity relies on LC–MS only. The direct MS m/z values for all ions are detected and their signal intensities at a particular time recorded. The signal intensity from electrospray ionization has been reported to highly correlate with ion concentration, therefore the relative peptide levels between samples can be determined directly from these peak intensities. Because of the large amount of data collected from these experiments, sensitive computer algorithms are required for automated ion peak alignment and comparison. Label-free relative quantitation by spectral counts entails comparing the sum of the MS/MS spectra from a given peptide across multiple samples, which has been shown to directly correlate with protein abundance. Besides relative quantitation, label-free methods can be used to determine the absolute concentration of proteins in a sample. One method entails determining the exponentially modified protein abundance index (emPAI), which estimates protein abundance based on the number of peptides detected and the number of theoretically observed tryptic peptides for each protein, is used to determine the approximate absolute protein abundance in large-scale proteomic analyses. Another method, absolute protein expression (APEX) is based on spectral counts and uses correction factors to make protein abundance proportional to the number of peptides observed.

Flourescence resonance energy transfer (FRET) is an important tool to study protein–protein interactions, protein-DNA interactions, protein conformational changes and other molecular dynamics quantification. It is used to detect protein binding, providing spatial and temporal information about the interaction. It is a type of Fluorescence Spectroscopy using two fluorescent dyes with overlapping emission and absorption spectra, which is used to indicate proximity of labeled molecules. This technique is useful for studying interactions of molecules and protein folding. It uses fluorescent energy transfer to visualize protein interactions. Fluorophores are fused to the proteins of interest and then bombarded with light at the excitation wavelength. The first fluorophore transfers some of the energy it absorbs from the light source to the second fluorophore, which in turn emits some of its energy into the environment where it is visible with the use of a fluorescent microscope.

Since the discovery of FRET by Theodor Forster in 1948, it has become useful tool in biology for three reasons: (1) FRET is really sensitive in the range of 100 Å and below, the scale at which transactions between biological macromolecules and complexes occur; (2) the instrumentation is extremely sensitive and is readily amenable to miniaturization, high throughput, and automation; and (3) cell permeable and genetically encodable FRET probes enable the real‐time quantitation of dynamic cellular processes in live cells.

Mass spectrometry (MS) is an important emerging method for the characterization of proteins. MS is a technique in which gas phase molecules are ionized and their mass-to-charge ratio is measured by observing acceleration differences of ions when an electric field is applied (Fig. 8.6). Lighter ions will accelerate faster and be detected first. If the mass is measured with precision then the composition of the molecule can be identified. In the case of proteins, the sequence can be identified. Most samples submitted to MS are a mixture of compounds. A spectrum is acquired to give the mass-to-charge ratio of all compounds in the sample. MS throws light on molecular mechanisms within cellular systems. It is used for identifying proteins, functional interactions, and it further allows for determination of subunits. Several configurations of mass spectrometers that combine electrospray ionization (ESI) and matrix-assisted laser desorption ionization (MALDI) with a variety of mass analyzers (linear quadrupole mass filter [Q], time-of-flight [ToF], quadrupole ion trap, and Fourier transform ion cyclotron resonance [FTICR] instrument) are routinely used (Yates et al. 2009).

Whole protein mass analysis is primarily conducted using either TOF, MS, or FTICR. These two instruments are preferable because of their wide mass range and in the case of FTICR, its high-mass accuracy. Mass analysis of proteolytic peptides is a much more popular method of protein characterization, as cheaper instrument designs can be used for characterization. Additionally, sample preparation is easier once whole proteins have been digested into smaller peptide fragments. The most widely used instrument for peptide mass analysis are the MALDI time-of-flight instruments as they permit the acquisition of peptide mass fingerprints (PMFs) at high pace. Multiple stage quadrupole-time-of-flight and the quadrupole ion trap also find use in this application.

The relative abundance of an ion can also be measured using MS. Different compounds have differential ionization capabilities, therefore intensity of an ion is not in a direct correlation to concentration. It is an analytical method which has a variety of uses outside of proteomics, such as isotope and dating, trace gas analysis, atomic location mapping, pollutant detection, and space exploration. This technique was discovered during studies of gas excitation in a charged environment, more than 100 years ago by J. J. Thomson in 1913.

The ionization methods used for the majority of biochemical analyses are:

ESI is one of the atmospheric pressure ionization (API) techniques and is well suited to the analysis of polar molecules ranging from less than 100 Da to more than 1,000,000 Da in molecular mass. In ESI the ions of interest are formed from solution by applying a high electric field to the tip of a capillary, from which the solution will pass through. The sample will be sprayed into the electric field along with a flow of nitrogen to promote desolvation. Droplets will form and will evaporate in a vacuumed area. This causes an increase in charge on the droplets and the ions are now said to be multiply charged. These multiply charged ions can then enter the analyzer (Andersen et al. 1996). ESI is a method of choice because of the following properties: (1) The “softness” of the phase conversion process allows very fragile molecules to be ionized intact and even in some non-covalent interactions to be preserved for MS analysis. (2) The eluting fractions through liquid chromatography can then be sprayed into the mass spectrometer, allowing for the further analysis of mixtures. (3) The production of multiply charged ions allow for the measurement of high-mass biopolymers. Multiple charges on the molecule will reduce its mass to charge ratio when compared to a single charged molecule. Multiple charges on a molecule also allow for improved fragmentation which in turn allows for a better determination of structure.

MALDI (Hillenkamp et al. 1991) deals with thermolabile, non-volatile organic compounds especially those of high molecular mass and is used successfully in biochemical areas for the analysis of proteins, peptides, glycoproteins, oligosaccharides, and oligonucleotides. It is relatively straightforward to use and reasonably tolerant to buffers and other additives. The mass accuracy depends on the type and performance of the analyzer of the mass spectrometer, but most modern instruments are capable of measuring masses to within 0.01 % of the molecular mass of the sample, at least up to ca. 40,000 Da. In MALDI, the molecular ions of interest are formed by pulses of laser light impacting on the sample isolated within an excess of matrix molecules. This enables the determination of masses of large biomolecules and synthetic polymers greater than 200,000 Daltons without degradation of the molecule of interest. The advantages of MALDI are its robustness, high speed, and relative immunity to contaminants and biochemical buffers. A type of mass spectrometer often used with MALDI is TOF or Time-of-Flight mass spectrometry. This enables fast and accurate molar mass determination along with sequencing repeated units and recognizing polymer additives and impurities. This technique is based on an ultraviolet absorbing matrix where the matrix and polymer are mixed together along with excess matrix and a solvent to prevent aggregation of the polymer. This mixture is then placed on the tip of a probe; then the solvent is removed while under vacuum conditions. This creates co-crystallized polymer molecules that are dispersed homogeneously within the matrix. A pulsing laser beam is set to an appropriate frequency and energy is shot to the matrix, which becomes partially vaporized. As a result the homogeneously dispersed polymer within the matrix is carried into the vapor phase and becomes charged.