Protein electrophoresis is a standard laboratory technique by which charged protein molecules are transported through a solvent by an electrical field. Both proteins and nucleic acids may be separated by electrophoresis, which is a simple, rapid, and sensitive analytical tool. Most biological molecules carry a net charge at any pH other than their isoelectric point and will migrate at a rate proportional to their charge density. The mobility of a molecule through an electric field will depend on the following factors: field strength, net charge on the molecule, size and shape of the molecule, ionic strength, and properties of the matrix through which the molecule migrates (e.g., viscosity, pore size). Polyacrylamide and agarose are two support matrices commonly used in electrophoresis. These matrices serve as porous media and behave like a molecular sieve. Agarose has a large pore size and is suitable for separating nucleic acids and large protein complexes. Polyacrylamide has a smaller pore size and is ideal for separating majority of proteins and smaller nucleic acids.
Several forms of polyacrylamide gel electrophoresis (PAGE) exist, and each form can provide different types of information about proteins of interest. Denaturing and reducing sodium dodecyl sulfate PAGE (SDS-PAGE) with a discontinuous buffer system is the most widely used electrophoresis technique and separates proteins primarily by mass. Nondenaturing PAGE, also called native-PAGE, separates proteins according to their mass/charge ratio. Two-dimensional (2D) PAGE separates proteins by native isoelectric point in the first dimension and by mass in the second dimension.
SDS-PAGE separates proteins primarily by mass because the ionic detergent SDS denatures and binds to proteins to make them uniformly negatively charged. Thus, when a current is applied, all SDS-bound proteins in a sample will migrate through the gel toward the positively charged electrode. Proteins with less mass travel more quickly through the gel than those with greater mass because of the sieving effect of the gel matrix. Once separated by electrophoresis, proteins can be detected in a gel with various stains, transferred onto a membrane for detection by western blotting and/or excised and extracted for analysis by mass spectrometry. Protein gel electrophoresis is, therefore, a fundamental step in many kinds of proteomics analysis.
Polyacrylamide is the material of choice for preparing electrophoretic gels to separate proteins by size. Polyacrylamide gels are prepared by mixing acrylamide with bisacrylamide to form a crosslinked polymer network when the polymerizing agent, ammonium persulfate (APS), is added. TEMED (N,N,N’,N'-tetramethylenediamine) catalyzes the polymerization reaction by promoting the production of free radicals by APS. At this stage it becomes polyacrylamide.
Polymerization and crosslinking of acrylamide. The ratio of bisacrylamide (N,N'-methylenediacrylamide) to acrylamide, as well as the total concentration of both components, affects the pore size and rigidity of the final gel matrix. These, in turn, affect the range of protein sizes (molecular weights) that can be resolved.
The size of the pores created in the gel is inversely related to the polyacrylamide percentage (concentration). For instance, a 7% polyacrylamide gel has larger pores than a 12% polyacrylamide gel. Low-percentage gels are used to resolve large proteins, and high-percentage gels are used to resolve small proteins. "Gradient gels" are specially prepared to have a low percentage of polyacrylamide at the top (beginning of sample path) and a high percentage at the bottom (end), enabling a broader range of protein sizes to be separated.
Electrophoresis gels are formulated in buffers that enable electrical current to flow through the matrix. The prepared solution is poured into the thin space between two glass or plastic plates that form a cassette. This process is referred to as casting a gel. Once the gel polymerizes, the cassette is mounted (usually vertically) into an apparatus so that the top and bottom edges are placed in contact with buffer chambers containing a cathode and an anode, respectively. The running buffer contains ions that conduct current through the gel. When proteins are loaded into wells at the top edge and current is applied, the proteins are drawn by the current through the matrix slab and separated by the sieving properties of the gel.
To obtain optimal resolution of proteins, a stacking gel is cast over the top of the resolving gel. The stacking gel has a lower concentration of acrylamide (e.g., 7% for larger pore size), lower pH (e.g., 6.8), and a different ionic content. This allows the proteins in a loaded sample to be concentrated into one tight band during the first few minutes of electrophoresis before entering the resolving portion of a gel. A stacking gel is not necessary when using a gradient gel, as the gradient itself performs this function.
Polyacrylamide gel electrophoresis in progress. Prepared gel cassettes are inserted into a gel tank, in this case the Invitrogen Mini Gel Tank, which holds two mini gels at a time. After wells are loaded with protein samples, the gels submerged in a conducting running buffer, and electrical current is applied, typically for 20 to 40 minutes. Run times vary according to the size and percentage of the gel and gel chemistry.
In SDS-PAGE, the gel is cast in a buffer containing sodium dodecyl sulfate (SDS), an anionic detergent. SDS denatures proteins by wrapping around the polypeptide backbone. By heating the protein sample between 70-100°C in the presence of excess SDS and thiol reagent, disulfide bonds are cleaved, and the protein is fully dissociated into its subunits. Under these conditions most polypeptides bind SDS in a constant weight ratio (1.4 g of SDS:1 g of polypeptide). The intrinsic charges of the polypeptide are insignificant compared to the negative charges provided by the bound detergent so that the SDS-polypeptide complexes have essentially the same negative charge and shape. Consequently, proteins migrate through the gel strictly according to polypeptide size with very little effect from compositional differences. The simplicity and speed of this method, plus the fact that only microgram quantities of protein are required, have made SDS-PAGE the most widely used method for determination of molecular mass in a polypeptide sample. Proteins from almost any source are readily solubilized by SDS so the method is generally applicable.
When a set of proteins of known mass are run alongside samples in the same gel, they provide a reference by which the mass of sample proteins can be determined. These sets of reference proteins are called mass markers or molecular weight markers (MW markers), protein ladders, or size standards, and they are available commercially in several forms.
In native-PAGE, proteins are separated according to the net charge, size, and shape of their native structure. Electrophoretic migration occurs because most proteins carry a net negative charge in alkaline running buffers. The higher the negative charge density (more charges per molecule mass), the faster a protein will migrate. At the same time, the frictional force of the gel matrix creates a sieving effect, regulating the movement of proteins according to their size and three-dimensional shape. Small proteins face only a small frictional force, while larger proteins face a larger frictional force. Thus native-PAGE separates proteins based upon both their charge and mass.
Because no denaturants are used in native-PAGE, subunit interactions within a multimeric protein are generally retained and information can be gained about the quaternary structure. In addition, some proteins retain their enzymatic activity (function) following separation by native-PAGE. Thus, this technique may be used for preparation of purified, active proteins.
Following electrophoresis, proteins can be recovered from a native gel by passive diffusion or electro-elution. To maintain the integrity of proteins during electrophoresis, it is important to keep the apparatus cool and minimize denaturation and proteolysis. pH extremes should generally be avoided in native-PAGE, as they may lead to irreversible damage, such as denaturation or aggregation, to proteins of interest.
The most common form of protein gel electrophoresis is comparative analysis of multiple samples by one-dimensional (1D) electrophoresis. Gel sizes range from 2 x 3 cm (tiny) to 15 x 18 cm (large format). The most popular size (approx. 8 x 8 cm) is usually referred to as a "mini gel". Medium-sized gels (8 x 13 cm) are called midi gels. Small gels require less time and reagents than their larger counterparts and are suited for rapid protein screening. However, larger gels provide better resolution and are needed for separating similar proteins or a large number of proteins.
Protein samples are added to sample wells at the top of the gel. When the electrical current is applied, the proteins move down through the gel matrix, creating what are called lanes of protein bands. Samples that are loaded in adjacent wells and electrophoresed together are easily compared to each other after staining or other detection strategies. The intensity of staining and thickness of protein bands are indicative of their relative abundance. The positions (height) of bands within their respective lanes indicate their relative sizes (and/or other factors affecting their rate of migration through the gel).
Protein lanes and bands in 1D SDS-PAGE. Depicted here is a protein ladder, purified proteins and E. coli lysate loaded on a 4–20% gradient Novex Tris-Glycine gel; Lanes 1, 5, 10: 5 µL Thermo Scientific PageRuler Unstained Protein Ladder); lanes 2, 6, 9: 5 µL Mark12 Unstained Standard; lane 3: 10 µg E. coli lysate (10 µL sample volume); lane 4: 6 µg BSA (10 µL sample volume); lane 7: 6 µg hIgG (10 µL sample volume); lane 8: 20 µg E. coli lysate (20 µL sample volume). Electrophoresis was performed using the Mini Gel Tank. Sharp, straight bands were observed after staining with SimplyBlue SafeStain. Images were acquired using a flatbed scanner.
Multiple components of a single sample can be resolved most completely by two-dimensional electrophoresis (2D-PAGE). The first dimension separates proteins according to their native isoelectric point (pI) using a form of electrophoresis called isoelectric focusing (IEF). The second dimension separates by mass using ordinary SDS-PAGE. 2D PAGE provides the highest resolution for protein analysis and is an important technique in proteomic research, where resolution of thousands of proteins on a single gel is sometimes necessary.
To perform IEF, a pH gradient is established in a tube or strip gel using a specially formulated buffer system or ampholyte mixture. Ready-made IEF strip gels (called immobilized pH gradient strips or IPG strips) and required instruments are available from certain manufacturers. During IEF, proteins migrate within the strip to become focused at the pH points at which their net charges are zero. These are their respective isoelectric points.
The IEF strip is then laid sideways across the top of an ordinary 1D gel, allowing the proteins to be separated in the second dimension according to size.
Example 2-D electrophoresis data. In the first dimension, one or more samples are resolved by isoelectric focusing (IEF) in strip gels. IEF is usually performed using precast immobilized pH-gradient (IPG) strips on a specialized horizontal electrophoresis platform. For the second dimension, a gel containing the pI-resolved sample is laid across to top of a slab gel so that the sample can then be further resolved by SDS-PAGE.
Three basic types of buffers are required: the gel casting buffer, the sample buffer, and the running buffer that fills the electrode reservoirs. Electrophoresis may be performed using continuous or discontinuous buffer systems. A continuous buffer system, which utilizes only one buffer in the gel, sample, and gel chamber reservoirs, is most often used for nucleic acid analysis and rarely used for protein gel electrophoresis. Proteins separated using a continuous buffer system tend to be diffuse and poorly resolved. Conversely, discontinuous buffer systems utilize a different gel buffer and running buffer. These systems also use two gel layers of different pore sizes and different buffer compositions (the stacking and separating gels). Electrophoresis using a discontinuous buffer system results in concentration of the sample and higher resolution. The various commonly used discontinuous gel buffer systems as summarized below.
The most widely used gel system for separating a broad range of proteins is the Laemmli system. The classical Laemmli system, consisting of Tris-glycine gels and Tris-glycine running buffer, can be used for both SDS-PAGE and native PAGE. This system is used widely because reagents for casting Tris-glycine gels are relatively inexpensive and readily available. Gels using this chemistry can be made in a variety gel formats and percentages.
The formulation of this discontinuous buffer system creates a stacking effect to produce sharp protein bands at the beginning of the electrophoretic run. A boundary is formed between chloride, the leading ion, and glycinate, the trailing ion. Tris buffer provides the common cations. As proteins migrate into the resolving gel, they are separated according to size. Tris-glycine gels are used in conjunction with Laemmli sample buffer, and Tris/glycine/SDS running buffer is used for denaturing SDS-PAGE. Native PAGE is performed using native sample and running buffers without denaturants or SDS. The pH and ionic strength of the buffer used for running the gel (Tris, pH 8.3) are different from those of the buffers used in the stacking gel (Tris, pH 6.8) and the resolving gel (Tris, pH 8.8). The highly alkaline operating pH of the Laemmli system may cause band distortion, loss of resolution, or artifact bands.
In contrast to conventional Tris-glycine gels, Bis-Tris HCI–buffered gels run closer to neutral pH, thus offering enhanced stability and greatly extended shelf-life over Tris-glycine gels (up to 16 months at room temperature). The neutral pH provides reduced protein degradation and is good for applications where high sensitivity is required such as analysis of posttranslational modifications, mass spectrometry, or sequencing.
For Bis-Tris gels, chloride serves as the leading ion and MES or MOPS act as the trailing ion. Bis-Tris buffer forms the common cation. Markedly different protein migration patterns are produced depending on whether a Bis-Tris gel is run with MES or MOPS denaturing running buffer: MES buffer is used for smaller proteins, and MOPS buffer is used for mid-sized proteins.
Due to differences in ionic composition and pH, gel patterns obtained with Bis-Tris gels cannot be compared to those obtained with Tris-glycine gels. To prevent protein reoxidation, Bis-Tris gels must be run with alternative reducing agents such as sodium bisulfite. Reducing agents frequently used with Tris-glycine gels, such as beta-mercaptoethanol and dithiothreitol (DTT), do not undergo ionization at low pH levels and are not able to migrate with proteins in a Bis-Tris gel.
Tris-acetate gel chemistry enables the optimal separation of high molecular weight proteins. Tris-acetate gels use a discontinuous buffer system involving three ions- acetate, tricine and tris. Acetate serves as a leading ion due to its high affinity to the anode relative to other anions in the system. Tricine serves as the trailing ion.Tris-acetate gels can be used with both SDS-PAGE and native PAGE running buffers. Compared with Tris-glycine gels, Tris-acetate gels have a lower pH, which enhances the stability of these gels and minimizes protein modifications, resulting in sharper bands.
The Tris-Tricine gel system is a modification of the Tris-glycine gel system and is optimized to resolve low molecular weight proteins in the range of 2–20 kDa. As a result of reformulating the Laemmli running buffer and using Tricine in place of glycine, SDS-polypeptides form behind the leading ion front rather than running with the SDS front, thus allowing for their separation into discrete bands.
Zymogram gels are Tris-glycine gels containing gelatin or casein and are used to characterize proteases that utilize them as substrates. Samples are run under denaturing conditions, but due to the absence of reducing agents, proteins undergo renaturation. Proteolytic proteins present in the sample consume the substrate, generating clear bands against a background stained blue.
The choice of whether to use one chemistry or another depends on the abundance of the protein separating, the size of the protein and the downstream application. For separation of a broad range of proteins two chemistries: Bis-Tris and Tris-glycine are well suited. Bis-Tris gel chemistry provides greater sensitivity for protein detection compared to Tris-glycine gel chemistry. Choose Bis-Tris gel chemistry when you have a low abundance of protein or when the downstream application requires high protein integrity, such as posttranslational modification analysis, mass spectrometry, or sequencing.
|Protein sample type||Broad range MW (6-400 kDa)||Broad range MW (6-400 kDa)||High range MW (40-500 kDa)||Low range MW (2.5-40 kDa)|
|Chemistry benefits||Neutral pH for high-sensitivity applications and reduced protein degradation||Traditional Laemmli-style||Analysis of high molecular weight proteins; neutral pH||Analysis of low molecular weight proteins|
|Recommended for||Western blotting, mass spectrometry, posttranslationally modified proteins, dilute samples, and low-abundance proteins||Western blotting, in-gel staining, samples containing detergents and high salt, native- PAGE applications||High molecular weight proteins, western blotting, mass spectrometry, posttranslationally modified proteins, native-PAGE applications||Low molecular weight proteins, western blotting, in-gel staining|
Protein samples prepared for SDS-PAGE analysis are denatured by heating in the presence of a sample buffer containing 1% SDS with or without a reducing agent such as 20mM DTT, 2-mercaptoethanol (BME) or Tris(2-carboxyethyl)phosphine (TCEP). The protein sample is mixed with the sample buffer and heated for 3 to 5 minutes (according to the specific protocol) then cooled to room temperature before it is pipetted into the sample well of a gel. Loading buffers also contain glycerol so that they are heavier than water and sink neatly to the bottom of the buffer-submerged well when added to a gel.
If a suitable, negatively charged, low-molecular weight dye is also included in the sample buffer, it will migrate at the buffer-front, enabling one to monitor the progress of electrophoresis. The most common tracking dyes for sample loading buffers are bromophenol blue, phenol red and Coomassie blue. The table below summarizes common sample buffers and running buffers used in the different gel buffer systems.
|Gel chemistry||Sample buffer||Running buffer||Selection criteria|
|Tris-glycine||Tris-glycine SDS sample buffer: Tris HCl (63 mM), glycerol (10%), SDS (2%), bromophenol blue (0.0025%), pH 6.8||Tris-glycine SDS: Tris base (25 mM), glycine (192 mM), SDS (0.1%), pH 8.3||Ease of preparation; relatively inexpensive, separation of broad range of molecular weight proteins|
|Bis-Tris||LDS sample buffer: Tris base (141 mM), Tris HCl (106 mM), LDS (2%), EDTA (0.51 mM), SERVA Blue G-250 (0.22 mM), phenol red (0.175 mM), pH 8.5||MES SDS: MES (50 mM), Tris base (50 mM), SDS (0.1%), EDTA (1 mM), pH 7.3|
MOPS SDS: MOPS (50 mM), Tris base (50 mM), SDS (0.1%), EDTA (1 mM), pH 7.7
|Relatively long shelf life; room temperature storage; neutral pH minimizes protein modifications, separation of broad range of molecular weight proteins|
|Tris-Acetate||LDS sample buffer: Tris base (141 mM), Tris HCl (106 mM), LDS (2%), EDTA (0.51 mM), SERVA Blue G-250 (0.22 mM), phenol red (0.175 mM), pH 8.5||Tris-acetate SDS: Tris base (50 mM), Tricine (50 mM), SDS (0.1%), pH 8.24||Superior separation of protein complexes and high MW proteins; relatively long shelf life|
|Tris-Tricine||Tricine SDS sample buffer: Tris HCl (450 mM), glycerol (12%), SDS (4%), Coomassie Blue G (0.00075%), phenol red (0.0025%), pH 8.45||Tricine-SDS: Tris base (100 mM), tricine (100 mM), SDS (0.1%), pH 8.3||Ideal for separating peptides and low molecular weight proteins|
|Tris-glycine||Native sample buffer: Tris HCl (100 mM), glycerol (10%), bromophenol blue (0.00025%), pH 8.6||Tris-Glycine Native buffer: Tris base (25 mM), glycine (192 mM), pH 8.3||Retention of native protein structure|
|Tris-acetate||Native sample buffer: Tris HCl (100 mM), glycerol (10%), bromophenol blue (0.00025%), pH 8.6||Tris-Glycine Native buffer: Tris base (25 mM), glycine (192 mM), pH 8.3||Superior separation of protein complexes and high MW proteins|
|IEF||IEF Sample Buffer pH 3-7: Lysine (40 mM), glycerol (15%)|
IEF Sample Buffer pH 3-10: Arginine (20 mM), Lysine (20 mM), glycerol (15%)
|IEF cathode buffer pH 3-7: Lysine (40 mM)|
IEF cathode buffer pH 3-10: Arginine (20 mM), lysine (20 mM)
IEF anode buffer: phosphoric acid 85% (7 mM)
|Use to separate proteins according to isoelectric point (pI) rather than molecular weight|
|Zymogram||Tris-glycine SDS: Tris HCl (63 mM), glycerol (10%), SDS (2%), bromophenol blue (0.0025%), pH 6.8||Tris-glycine SDS: Tris base (25 mM), glycine (192 mM), SDS (0.1%), pH 8.3||Gelatin or casein gels provide substrates used to detect proteases|
|Gel Type||Voltage||Expected current||Run time|
|Tris-glycine||Denaturing: 125 volts constant|
Native: 20-125 volts constant
|Denaturing: 30-40 mA (start), 8-12 mA (end)|
Native: 6-12 mA (start), 3-6 mA (end)
|Denaturing: 90 min|
Native: 1-12 hr
|Bis-Tris||200 volts constant||Non-reducing: 100-125 mA (start), 60-70 mA (end) Reducing: 110-125 mA (start), 70-80 mA (end)||35-50 min|
|Tris-Acetate||Denaturing: 150 volts constant|
Native: 20-150 volts constant
|Denaturing and Native:|
40-55 mA (start), 25-40 mA (end)
|Denaturing: 60 min|
Native: 1-12 hr
|Tricine||125 volts constant||80 mA (start), 40 mA (end)||90 min|
|IEF||100 volts for 1hr, 200 volts for 1hr, 500 volts for 30 min||5 mA (start), 6 mA (end)||2.5 hr|
|Zymogram||125 volts constant||30-40 mA (start), 8-12 mA (end)||90 min|
Traditionally, researchers casted their own gels using standard recipes that are widely available in protein methods literature. More laboratories are moving to the convenience and consistency afforded by commercially available, ready-to-use precast gels. Precast gels are available in a variety of percentages, including difficult-to-pour gradient gels that provide excellent resolution and that separate proteins over the widest possible range of molecular weights. Precast gels are also available in the different buffer formulations (e.g., Tris-glycine, Bis-Tris, Tris-acetate, Tricine), which are designed to optimize shelf life, run time, and/or protein resolution.
For researchers who require unique gel formulations not available as precast gels, a wide range of reagents and equipment are available for pouring gels. However, technological innovations in buffers and gel polymerization methods enable manufacturers to produce gels with greater uniformity and longer shelf life than individual researchers can prepare on their own with traditional equipment and methods. In addition, precast polyacrylamide gels eliminate the need to work with the acrylamide monomer, which is a known neurotoxin and suspected carcinogen.
Precast vs. handcast protein gels for SDS-PAGE. Polyacrylamide gels can be purchased precast and ready- to- use (left) or prepared from reagents in the lab using a gel-casting system (right). Pictured here are the Novex Tris-Glycine Mini Gels, WedgeWell format (left) and the SureCast Gel Handcast System.
To perform protein gel electrophoresis, the polyacrylamide gel and buffer must be placed in an electrophoresis chamber that is connected to a power source, and which is designed to conduct current through the buffer solution. When current is applied, the smaller molecules migrate more rapidly and the larger molecules migrate more slowly through the gel matrix. Multiple gel chamber designs exist. The choice of equipment is usually based on these factors: the dimensions of the gel cassette, with some tank designs accommodating more cassette sizes than others; the nature of the protein target, and corresponding gel resolution requirements; and whether a precast or handcast gel, and vertical or horizontal electrophoresis system, has been selected.
Mini gel tank for protein gel electrophoresis. This gel tank holds up to two mini gels and is compatible with the Invitrogen SureCast Gel Handcast System, and with all Invitrogen precast gels. The unique tank design enables side-by-side gel loading and enhanced viewing during use.
To assess the molecular masses (sizes) of proteins in a gel, a prepared mixture containing several proteins of known molecular masses is run alongside the test sample in one or more lanes of the gel. Such sets of known proteins are called protein molecular weight (or mass) markers or protein ladders. A standard curve can be constructed from the distances migrated by each marker protein. The distance migrated by the unknown protein is then plotted, and the molecular weight is extrapolated from the standard curve.
Several kinds of ready-to-use protein molecular weight (MW) markers are available that are either unlabeled or prestained for different modes of detection. These are pre-reduced and, therefore, primarily suited for SDS-PAGE rather than native PAGE. MW markers can also be made detectable via specialized labels, such as a fluorescent tag, and by other methods.
Generally, protein mobility in SDS gels is a function of the length of the protein in its fully denatured state. By constructing a standard curve with protein standards of known molecular weights, the molecular weight of a sample protein can be calculated based upon its relative mobility. However, the same molecular weight standard may have slightly different mobility and therefore, different apparent molecular weight when run in different SDS-PAGE buffer systems.
When using SDS-PAGE for molecular weight calibration, slight deviations from the true molecular weight of a protein (definitively calculated from the known amino acid sequence) can occur mostly because of the retention of varying degrees of secondary structure in the protein, even in the presence of SDS. This phenomenon is more prevalent in proteins with highly organized secondary structures (such as collagens, histones, or highly hydrophobic membrane proteins) and in peptides, where the effect of local secondary structure becomes magnified relative to the total size of the peptide.
It has also been observed that slight differences in protein mobilities occur when the same proteins are run in different SDS-PAGE buffer systems. Each SDS-PAGE buffer system has a different pH, which affects the charge of a protein and its binding capacity for SDS. The degree of change in protein mobility is usually small in natural proteins but is more pronounced with atypical or chemically modified proteins, such as pre-stained standards. Apparent molecular weight values for pre-stained standards will vary between gel systems- it is important to use the apparent molecular weights that matches your gel for the most accurate calibration of your sample proteins.
For Research Use Only. Not for use in diagnostic procedures.