Once protein bands have been separated by electrophoresis, they can be visualized using different methods of in-gel detection, each with particular advantages and disadvantages. Over the past several decades, demand for improved sensitivity for small sample sizes and compatibility with downstream applications and detection instrumentation have driven the development of several basic staining methods. Here we discuss the general principles of protein gel staining and describe several staining methods.
General principles of gel staining
The first step after performing denaturing polyacrylamide gel electrophoresis (SDS-PAGE) is to disassemble the gel cassette and place the thin polyacrylamide gel in a tray filled with water or buffer. The electrophoresed proteins exist as concentrated "bands" embedded within each lane of the porous polyacrylamide gel matrix. Typically, the proteins are still bound to the anionic detergent (SDS), and the entire gel matrix is saturated in running buffer.
To make the proteins visible, a protein-specific, dye-binding or color-producing chemical reaction must be performed on the proteins within the gel. Depending on the particular chemistry of the stain, various steps are necessary to retain, or fix, the proteins in the gel matrix and to facilitate the necessary chemical reaction. All steps are done in solution, i.e., with the gel suspended in a tray filled with one liquid reagent or another.
Given the common constraints of this format, most staining methods involve some version of the same general incubation steps:
- A water wash to remove electrophoresis buffers from the gel matrix
- An acid or alcohol wash to condition or fix the gel to limit diffusion of protein bands from the matrix
- Treatment with the staining reagent to allow the dye or chemical to diffuse into the gel and bind to (or react with) the proteins
- Destaining to remove excess dye from the gel matrix background
Depending on the particular staining method, two or more of these functions can be accomplished with one step. For example, a dye reagent that is formulated in an acidic buffer can effectively fix and stain in one step. Conversely, certain functions require several steps. For example, silver staining requires both a staining reagent step and a developer step to produce the colored reaction product.
Protein Gel Electrophoresis Technical Handbook
This 88-page handbook provides detailed description about all aspects of protein electrophoresis from sample and gel preparation to choice of molecular weight markers. In addition, it contains extensive information about our portfolio of high-quality protein electrophoresis products including gels, stains, molecular weight markers, running buffers, and blotting products for your experiments.
The most common method of in-gel protein detection is staining with Coomassie dye. Several recipes for Coomassie staining reagents exist in the literature and use either the G-250 (“colloidal”) or R-250 form of the dye. Colloidal Coomassie can be formulated to effectively stain proteins within 1 hour and requires only water (no methanol or acetic acid) for destaining.
In acidic conditions, Coomassie dye binds to basic and hydrophobic residues of proteins, changing in color from a dull reddish-brown to intense blue (see previous images on this page). As with all staining methods, Coomassie staining detects some proteins better than others, based on the chemistry of action and differences in protein composition. Thus, Coomassie staining can detect as little as 8–10 ng per band for some proteins and 25 ng per band for most proteins.
Gel staining with Coomassie dye. Two-fold dilutions of protein extracts were run on an Invitrogen™ NativePAGE™ 3–12% Bis-Tris Protein Gel using a Mini Gel Tank. Following electrophoresis, the gel was stained with Coomassie dye and imaged using a flatbed scanner. Lanes 1 and 10: blank; lanes 2 and 6: 5 μL Invitrogen™ NativeMark™ Unstained Protein Standard; lanes 3, 4 and 5: 10, 5, and 2.5 μg spinach chloroplast extract; lanes 7, 8, and 9: 10, 5, and 2.5 μg bovine mitochondrial extract.
Coomassie dye staining is especially convenient because it involves a single ready-to-use reagent and does not permanently chemically modify the target proteins. An initial water wash step is necessary to remove residual SDS, which interferes with dye binding. Then the staining reagent is added, usually for about 1 hour; finally, a water or simple methanol:acetic acid destaining step is used to wash away excess unbound dye from the gel matrix. Because no chemical modification occurs, excised protein bands can be completely destained and the proteins recovered for analysis by mass spectrometry or sequencing.
Coomassie staining and other traditional staining methods require several long incubation and wash steps. To expedite the staining process, more rapid staining protocols have been developed using powered (electrophoretic) devices such as the Thermo Scientific™ Pierce™ Power Stainer.
The Thermo Scientific™ Pierce™ Power Stainer. This powered device enables rapid (6–11 min) Coomassie dye staining of proteins in polyacrylamide gels, including the removal of unbound stain, in a single step. The small, easy-to-use device consists of the Pierce™ Power Station and Pierce™ Power Stain Cassette, which accommodates up to two mini gels or one midi gel at a time. The staining procedure is designed exclusively for use with Pierce™ Power Staining Kits.
Silver staining is the most sensitive colorimetric method for detecting total protein. The technique involves the deposition of metallic silver onto the surface of a gel at the locations of protein bands. Silver ions (from silver nitrate in the staining reagent) interact and bind with certain protein functional groups. The strongest interactions occur with carboxylic acid groups (Asp and Glu), imidazole (His), sulfhydryls (Cys), and amines (Lys). Various sensitizer and enhancer reagents are essential for controlling the specificity and efficiency of silver ion binding to proteins and effective conversion (development) of the bound silver to metallic silver. The development process is essentially the same as for photographic film: silver ions are reduced to metallic silver, resulting in a brown-black color.
Gel staining with silver stain. Samples were separated on an Invitrogen™ NuPAGE™ 4–12% Bis-Tris Protein Gel and stained with the Invitrogen™ SilverXpress™ Kit. Lanes 1–5: Invitrogen™ Mark12™ Unstained Standard (blend of 12 purified proteins), serial 2-fold dilutions ranging from 1:4 to 1:64; lane 6: 1.6 ng BSA; lane 7: 0.8 ng BSA; lane 8: E. coli lysate diluted 1:20; lane 9: E. coli lysate diluted 1:80; lane 10: replicate of lane 1.
Silver staining protocols require several steps, which are affected by reagent quality as well as incubation times and thickness of the gel. An advantage of commercially available silver staining kits is that the formulations and protocols are optimized and consistently manufactured, helping to maximize consistency of results from experiment to experiment. Kits with optimized protocols are robust and easy to use, detecting less than 0.5 ng of protein in typical gels.
Silver stains use either glutaraldehyde or formaldehyde as the enhancer. These reagents can cause chemical crosslinking of the proteins in the gel matrix, limiting compatibility with destaining and elution methods for analysis by mass spectrometry (MS). Therefore, optimization of sensitivity vs. protein recoverability is critical when employing silver staining as part of an MS workflow.
Silver stain formulations can be made such that protein bands stain black, blue-brown, red, or yellow, depending on their charge and other characteristics. This is particularly useful for differentiating overlapping spots on 2D gels.
Zinc staining is unlike all other staining methods. Instead of staining the proteins, this procedure stains all areas of the polyacrylamide gel in which there are no proteins. Zinc ions complex with imidazole, which precipitates in the gel matrix except where SDS-saturated proteins are located. The milky-white precipitate renders the background opaque while the protein bands remain clear. The process is short (about 15 minutes), and the gel can be photographed by viewing it over a dark background. Zinc staining is as sensitive as typical silver staining (detects less than 1 ng of protein), and no fixation steps are required. Furthermore, the stain is easily removed, making this method compatible with MS or western blotting.
Gel staining with zinc stain. A 2-fold dilution series of a protein mixture was separated by protein gel electrophoresis using a 15-well mini gel. Subsequently the gel was stained using the Thermo Scientific™ Pierce™ Zinc Reversible Stain Kit, and then photographed with the gel placed over a dark blue background. The sensitivity on this gel is 0.25 ng, as indicated by the bands that are visible in the last lane.
Recent improvements in fluorescence imaging instruments and fluorescent applications have resulted in greater demand for fluorescent stains. A number of fluorescent stains for total protein have been introduced in recent years. Newer fluorescent total-protein stains provide exceptional fluorescent staining performance with fast and easy procedures. The most useful are those whose excitation and emission maxima correspond to common filter sets and laser settings of popular fluorescence imaging instruments.
Protein gel stained with fluorescent dyes. 2D gel stained with Invitrogen™ SYPRO™ Ruby protein gel stain and Invitrogen™ Pro-Q™ Emerald 300 reagent. Cohn fractions II and III from cow plasma were combined and resolved on a 2D gel. The gel was first stained with Pro-Q Emerald 300 reagent (left), followed by staining with the SYPRO Ruby stain (right).
Most fluorescent stains involve simple dye-binding mechanisms rather than chemical reactions that alter protein functional groups. Therefore, most are compatible with destaining and protein recovery methods for downstream analysis by MS. Accordingly, these stains are frequently used in both 1D and 2D applications.
Functional group–specific stains
Sometimes it is desirable to detect a subset of proteins rather than all of the proteins in a sample. Glycoproteins and phosphoproteins are classified as such on the basis of a particular chemical moiety (i.e., polysaccharides and phosphate groups, respectively). When a dye-binding or color-producing chemistry can be designed to detect one of these functional groups, it can be used as the basis for a specific gel stain.
Phosphoprotein and total protein visualization in a 2D gel. Protein lysates obtained from a Jurkat T-cell lymphoma line were separated by 2D gel electrophoresis and subsequently stained with Invitrogen™ Pro-Q™ Diamond phosphoprotein gel stain (blue) followed by SYPRO Ruby protein gel stain (red). The gel was dried and imaged on an FLA-3000 scanner (Fuji). Shown is a digitally pseudocolored composite overlaid image.
Proteins that have been post-translationally modified by glycosylation can be detected by a procedure that involves chemical activation of the carbohydrate into a reactive group. The method works by fixing the proteins in the gel and then oxidizing the sugar residues with sodium meta-periodate. The resulting aldehyde groups can then be reacted with an amine-containing dye. In older literature, this method is known as the periodate acid–Schiff (PAS) technique. A subsequent reduction step stabilizes the dye–protein bond. Both colorimetric and fluorescent dyes have been used for this technique, and glycoprotein stain kits are available commercially.
Various protein gel staining methods, both colorimetric and fluorescent, have also been developed to detect phosphorylated proteins and His-tagged fusion proteins. For instance, certain gel stains selectively stain phosphoproteins and His-tags in acrylamide gels, without the need for blotting or phosphoprotein-specific or His-tag–specific antibodies and western blot analysis.
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