Western blotting part I: Protein isolation (3' 27'')

Western blotting part II: Polyacrilamide-gel electrphoresis (6' 25'')

Western blotting part III: transfer, blotting and visualization (4' 23'')

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Separating proteins with SDS-PAGE

https://www-jove-com.bibliopass.unito.it/v/5058/separating-protein-with-sds-page

SDS-PAGE is a technique used by many researchers to separate mixtures of proteins by size. Successful completion of this technique is an essential first step for many methods of protein analysis, like immunoblotting. By itself, it is a useful tool in assessing protein size and purity.

In order to understand the SDS-PAGE technique, you must first understand its principle components. SDS-PAGE stands for Sodium Dodecyl Sulfate Poly-Acrylamide Gel Electrophoresis. Sodium-Dodecyl Sulfate, the first part of this, or “SDS”, is an anionic detergent. This means that it is composed of a hydrophilic group with a net negative charge and a long hydrophobic chain with neutral charge.

The hydrophobic chain blankets proteins in proportion to their mass at a rate of 1.4 grams of SDS per gram of protein. This provides the proteins with the driving force necessary for size driven separation in an electric field.

Poly-Acrylamide Gel Electrophoresis utilizes a hydrogel made from polyacrylamide. Polyacrylamide is a polymer that forms a very regular matrix through which proteins can move. The more concentrated the gel is, the slower the proteins will traverse across it when exposed to an electric field. The process of using a spatially uniform electric field to influence an objects motion is known as electrophoresis.

SDS-PAGE is performed on proteins isolated from many different sources including cells in culture, tissues, blood, urine, and yeast.

Proteins from these various sources must first be separated from other cellular components using techniques including homogenization and centrifugation, often followed by the use of lysis buffers.

Once the protein is isolated, its concentration is often measured, to ensure equal loading of samples, by comparing the amount of protein in the sample to albumin standards in a Bicinchoninic acid, or BCA, colorimetric assay.

An important step to remember is the addition of the loading buffer. The loading buffer has 3 main functions. First, thanks to SDS and additional reducing agents, it denatures the proteins, which basically means it turns complex protein structures into a linear chain of amino acids. Secondly, it contains glycerin, which ensures that the sample doesn’t float away when it is loaded in the wells of the gel. And finally, most commercial loading buffers include a dye, such as bromophenol blue, which can be tracked to measure the progress of the electrophoresis step.

After adding the loading buffer, the samples need to be mixed and then boiled for 5 minutes. This allows the strong di-sulfide bonds in the proteins to be broken with the help of a reducing agent such as beta-mercaptoethanol. Once all the disulfide bonds are broken SDS can more evenly coat the proteins.

Next, the samples are quickly spun down and are then ready to be loaded into the gel system for electrophoresis.

Before the samples can be loaded, the gel system must first be assembled. This begins with the purchase or fabrication of a polyacrylamide gel. Premade gels are becoming more and more popular because acrylimide is neurotoxic and can cause brain damage. The gel cassette contains wells that are used to load the samples.

Once the gels are secured into place, the inner and outer chambers are filled with a buffer that contains the same concentration of ions used to make the gels. This creates an electrical circuit that passes seamlessly from the cathode, through the gel and into the anode.

Next, molecular weight ladders are typically loaded into the gel, followed by the samples. As the gel runs, the ladder will spread and create visible protein bands of known sizes. At the final steps, these bands can be used to calculate each protein’s size.

Once all the samples are loaded, the positive and negative terminals on the gel box are connected to a power source capable of maintaining a constant voltage for a long period of time. Gels are typically started at around 60V until the entire sample has entered the gel region called the “stacking gel”. Then, the voltage is increased to around 200V for 30 minutes to 1 hour depending on the size and concentration of the gel and the size of the protein of interest.

When electrophoresis is complete, the cassette is removed and opened to expose the gel. The gel can then be stained with a typical protein stain, such as coomassie blue or silver stain, to visualize the protein bands within the gel. Coomassie stain can detect bands with as little as 50 nanograms of protein, whereas silver stain can detect bands with as little as 1 nanogram of protein.

In two-dimensional gel electrophoresis, samples are separated by two separate properties on gels, one dimension at a time. First, samples are loaded and arranged according to their isoelectric points on localized pH gradient strips. The proteins in the strip are then denatured and are placed on top of a typical polyacrylamide gel where they are secured in place with fresh gel solution. Then, second dimension separation is performed by SDS-PAGE.

While isoelectric focusing isn’t the only option for 2-D gel electrophoresis, it is the most common. Two-dimensional gel electrophoresis is an invaluable tool that provides insights into protein complexes and sub-organelle organization.

After running the gel, a very common next step is to transfer the proteins from the gel onto a membrane made of PVDF or nylon for analysis. You can then use specific antibodies to probe the membrane and look for proteins of interest. Shown here are typical immunoblot results where the researcher used 3 different signal amplification techniques to determine which best shows the presence of the Pit-1 protein throughout a number of serial dilutions.

The western blot

Western Blotting is a powerful technique utilized by many researchers to identify the presence of specific proteins in an electrophoretically-separated sample using antibodies.

There are 3 principal stages of this technique that are essential for a quality outcome: Electroblotting, Immunoblotting, and Detection. Before these stages are attempted, SDS-PAGE, in which denatured proteins are separated by size in a polyacrylamide gel, must be performed.

Electroblotting, is also known as the Western “transfer” and requires a transfer cassette for holding together the “sandwich” as well as an apparatus for transferring protein from an acrylimide gel to a thin membrane. The electroblotting sandwich consists of the gel and a specialized membrane, sandwiched between two pieces of filter paper. During the transfer, an electric field is used, to move the proteins through the gel, where they become trapped on a membrane due to charged and hydrophobic interactions.

Immunoblotting uses antibodies to “probe” the membrane for specific proteins. Antibodies are large Y-shaped proteins that contain two fragments, also known as Fab regions, which bind to other proteins. The Fab region defines the specific epitope, or specific portion of a protein, to which an antibody will bind.

Monoclonal antibodies are antibodies that recognize a single epitope and are the preferred antibody type used for immunoblotting due to their specificity. In contrast, polyclonal antibodies are a series of different antibodies that target many epitopes on the same antigen – or protein for which an antibody has specificity. Monoclonal antibodies that recognize a linear epitope are preferred as that ensures the epitope can be found on a denatured, or linearized, protein. This is important because many antibodies only recognize conformational epitopes, which means that they recognize proteins in their native 3D state only.

In addition to Fab fragments, antibodies contain an Fc region, which is specific to the animal that produced the antibody. In immunoblotting, this region is mainly utilized as the epitope for a secondary antibody – an antibody which recognizes the first antibody that has bound to the protein you’re trying to detect.

In order to produce an observable signal, antibodies are often linked, through their Fc region, to a reporter enzyme, such as alkaline phosphatase or horseradish peroxidase. These reporter enzymes produce signals by reacting with substrates to cause color changes or produce light changes.

These changes can then be quantified using densitometry. Densitometry is the technique used to measure the density of a protein band using image analysis software to calculate the density of each band. The bands can then be directly quantified using reference standards or internally controlled using a control sample.

For a successful Western transfer, the gel is first equilibrated in transfer buffer for 15 minutes. The membrane should also be equilibrated according to manufactures instructions.

Next, the gel transfer sandwich is carefully prepared in transfer buffer to prevent bubbles from being trapped within the system. Bubbles trapped in the sandwich can be forced out by rolling over them with a small pipette. Even small bubbles can interrupt the transfer of proteins causing incomplete transfer as can be seen on the lower parts of this membrane.

Once assembled, additional transfer buffer is poured into the transfer chamber and it is run at 20-30 mA for 2-3 hours. Transfer buffer contains methanol, which enhances the binding of the proteins to the membrane.

Once the transfer is complete the cassette is disassembled and the quality of the transfer can be checked using a non-specific protein stain such as Ponceau S. This offers a quick and reversible detection of protein bands.

The first step of immunoblotting is to cover the remaining areas of the membrane with a dilute solution of proteins. This step is called blocking and is performed, in order to block the membrane and reduce the nonspecific binding of the antibody to the membrane. Typically, the blocking solution consists of either bovine serum albumin or non-fat dry milk dissolved in a buffered saline solution. This step is usually done for 1 hour to overnight on a shaker.

The next step is to incubate the membrane with the primary antibody diluted in the blocking solution. This step can take from 30 minutes to overnight and should be performed with gentle agitation.

Then, the membrane is thoroughly rinsed to reduce nonspecific background staining and the secondary antibody is added in blocking buffer to the membrane. After a short incubation, the membrane is again thoroughly rinsed.

Secondary antibodies are detectable thanks to the reporter enzymes that are tethered to them. Upon addition of the correct substrate, colorimetric or chemiluminescent changes can be imaged and then measured using densitometry techniques. In addition, the protein ladder provides a valuable size reference so that each protein’s linear size can be estimated based on how far it has migrated in the gel relative to the size markers in the ladder. Observing the size of proteins detected via immunoblotting is a good way to verify that the antibody is recognizing the correct protein and whether it is a monomer or a multiple copies of the protein that are being seen.

There are thousands of commercially available primary antibodies, which allows for the detection and quantification of specific proteins within a sample. Here a researcher uses an antibody for HIF-1α to measure the amount of this oxygen responsive transcription factor in cell cultures to screen for hypoxia. They also used an antibody for beta-actin to act as a loading control. As you can see, the cells cultured in normal oxygen conditions produced far less HIF-1 than those cultured in low oxygen conditions.

The combination of 2D gel electrophoresis and immunoblotting can provide valuable information into the presence of protein complexes. Here, samples were first run by protein complex size, and then denatured so each individual protein in the complex could be separated by their individual size. Antibodies for three different proteins show three unique complexes containing different numbers of these proteins with each complex arranged in a vertical line. The left most column contains beta-2 and MCP-21, as well as another protein not shown by these markers. The center column shows a complex that contained all 3 proteins, and the right column contained only beta-2 and MCP-21.

Immunoblotting can also be used to visualize protein-protein interactions. This is performed by first transferring proteins from a gel onto a nitrocellulose membrane and then probing that membrane with additional proteins. Immunoblotting is then used to detect the if the added proteins complexed with any of the proteins that were immobilized on the membrane.