Why beta mercaptoethanol western blots

Large proteins such as membrane receptors do not blot well, and overall transfer efficiency is lower. Wet-transfer shines in its ability to yield high efficiency across a wide range of protein sizes, thus offering the most flexibility.

When Drs. Burnette and Towbin published their seminal studies; electrophoretic transfer was carried out on nitrocellulose membranes. They remained the gold standard until the advent of polyvinylidene difluoride PVDF membranes.

Concisely, PVDF membranes outcompete nitrocellulose membranes in their protein binding capacity, chemical resistance, and enhanced transfer efficiency in the presence of SDS.

PVDF's higher adsorption of proteins and its chemical resistance allows for stripping and reprobing of membranes. Also, by inserting a small percentage of SDS in the transfer buffer, transfer efficiency markedly improves. However, noted protein sensitivity from PVDF could also increase background signal for analysis. Methanol in transfer buffer can shrink nitrocellulose membranes and precipitate out large proteins. Both types of membranes come in different pore sizes, and membrane pore size is directly related to protein weight.

Smaller proteins require smaller pore sizes, although a pore size of 0. Recent years have seen the development of unique membranes such as those used for near-infrared detection systems. As such, the type of membrane chosen should reflect the target protein and downstream detection systems.

Upon completing the electrophoretic transfer, proteins are now on the membrane, and two antibodies serve for probing and analysis.

The primary antibody that binds a specific region on the target protein is used to detect its presence on the membrane. The secondary antibody conjugates with a component used for analysis. This antibody indirectly binds the target protein by binding to the constant regions of the primary antibody Figure 1c. Since membranes have a high affinity for protein, before probing, membranes are incubated in a buffer to coat the remaining surface area.

Typically, blocking buffer proteins include either casein from powdered milk or bovine serum albumin BSA. Although casein is cheaper and suitable for most proteins, BSA is considered a better choice when the target protein is phosphorylated.

There is cross-reactivity from casein and phosphorylation-specific primary antibodies. Tween 20 is a nonionic detergent that helps remove peripherally bound proteins on the membrane. Probing both primary and secondary antibodies is done by incubating the membrane in a probing buffer of either the primary or secondary antibody in TBS-T. The membrane is first incubated in the primary probing buffer typically overnight in a cold room, and washed again with TBS-T.

The membrane is then incubated with the secondary probing buffer for about an hour and then washed as well. These washing steps are crucial to reduce background noise in the analysis. After probing and washing, the membrane is ready to be read. As mentioned earlier, the secondary antibody conjugates with a component-specific to the type of analysis.

Autoradiography was a common way to visualize bands but has declined in its popularity due to hazards associated with this method. It uses a radiolabeled isotope conjugated to the secondary antibody. More commonly, a chemiluminescence method is used. This method uses substrates that react with an enzyme-conjugated secondary antibody. The enzyme-mediated reaction produces light that is then recorded with an imaging system. More recently, secondary antibodies have been conjugated with fluorophores that are capable of being detected without the need for substrates.

This fluorescence-based detection is gaining popularity due to its capability of probing two target proteins via secondary antibodies with different wavelength fluorophores; this is a selective advantage for relative protein expression analysis as housekeeping proteins are visible alongside a protein of interest. The visualization of bands can serve different analytical purposes. Simply, the presence of bands can verify the expression of a protein, whereas the density of bands can show comparative relative protein expression.

A housekeeping protein is also probed to evaluate relative protein expression. A housekeeping protein is a ubiquitous protein that constitutively expresses in all cells. By normalizing the band densities of the target protein with those of the housekeeping protein, a statistically significant difference between sample types can be measured.

As highlighted earlier, a western blot has a considerable amount of steps. This lengthy process drives up the time and cost needed for accurate results. Western blotting is used to detect anti-HIV antibodies in human serum and urine samples. The protein samples from a known HIV-infected individual get separated by electrophoresis and then blotted on the nitrocellulose membrane.

Then a specific antibody is affixed to detect the protein. More recently, in commercial HIV western blot kits, viral proteins come affixed to the membrane. Antibodies from human urine or serum samples bind to these proteins, and anti-HIV antibodies are used to detect bands alongside quality controls. The western blot is also useful in detecting Lyme disease and atypical and typical bovine spongiform encephalopathy. Like any experiment, quality controls should be used to validate findings.

In a western blot, a positive control, negative control, loading control, and a no first-degree A-B control are all effective in achieving and maintaining robust experiments.

Controls are dedicated lanes wherein the sample is altered specifically for the control type. A positive control is a sample known to contain the target protein, whereas a negative control is known not to contain the target protein. This can be as general as different organ types or as specific as different cellular localization. For example, if an analysis of the expression of a nuclear protein is the aim, and subcellular fractioning is done to isolate this region, a negative control evaluates the quality of fractioning, non-specific binding of antibodies, and a false-positive.

Positive controls are powerful in verifying that the workflow is well-optimized even in the absence of bands in sample lanes. Also, a positive control can verify a negative result.

Loading control is a housekeeping protein such as alpha-tubulin or beta-actin. Probing with antibodies specific for a housekeeping protein checks for an equal amount of proteins per sample. A nonspecific secondary antibody can yield false positives. The specificity of a secondary antibody is evaluated by not incubating a membrane strip with the primary antibody. Unfortunately, there are many error arms in this method due to many steps and a lengthy workflow. Discussing every error, its cause, and the solution is outside the scope of this review.

The most common issues and their troubleshooting will be of focus below. When bands are not migrating equally down the gel, this pattern can exaggerate to a smiley pattern. This indicates that the gel has air bubbles, voltage is too high, or the volume of the loading sample is too large. Air bubbles within the gel can distort the migration of bands.

A constant voltage during electrophoresis is directly proportional to resistance, and since resistance and temperature are directly linked, a high voltage increases the temperature in the electrophoresis tank. Heat pockets and an overall increase in the temperature of running buffer can also alter migration. Before the buffer can warm up, a high voltage at the start of electrophoresis will rush bands and cause nonlinear migration.

A large volume of loading samples can cause spillover into other lanes, and these large bands can skew into another lane. If the detection system shows no signal across all lanes except the ladder, there are a multitude of possible causes.

It is best to first localize wherein the workflow that the error occurred. Typically, the most common culprits are poor transfer efficiency or poor probing.

By staining the membrane with Ponceau S, a membrane-safe red dye, bands can be visualized. If bands are well illustrated on the membrane, particularly in the area where target protein is expected to be, it indicates that transfer efficiency is not likely the cause.

If there are no bands, transfer settings must be altered. A washout of proteins can occur in which the proteins from the membrane migrate to the filter paper. This is due to transfer time being too high; reducing voltage and or transfer time can prevent washout. A poor transfer can also occur if little to no proteins were adsorbed on the membrane. The 2X is to be mixed in ratio with the sample.

SDS binds to proteins fairly specifically in a mass ratio of 1. In doing so, SDS confers a negative charge to the polypeptide in proportion to its length. Denatured polypeptides become rods of negative charge with equal charge densities per unit length. Therefore, migration is determined by molecular weight, rather than by the intrinsic charge of the polypeptide.

SDS grade is important for high-quality protein separation: a protein stained background along individual gel tracts with indistinct or slightly distinct protein bands are indicative of old or poor quality SDS. Inclusion of 2-mercaptoethanol or dithiothreitol in the buffer reduces disulphide bridges, which is necessary for separation by size. Glycerol is added to the loading buffer to increase the density of the sample to be loaded and hence maintain the sample at the bottom of the well, restricting overflow and uneven gel loading.

To visualize the migration of proteins it is common to include a small anionic dye molecule in the loading buffer eg bromophenol blue. Since the dye is anionic and small, it will migrate the fastest of any component in the mixture to be separated and provide a migration front to monitor the separation progress.

During protein sample treatment the sample should be mixed by vortexing before and after the heating step for best resolution. Alternatively, an antibody may recognize an epitope made up of non-contiguous amino acids. Although the amino acids of the epitope are separated from one another in the primary sequence, they are close to each other in the folded three-dimensional structure of the protein, and the antibody will only recognize the epitope as it exists on the surface of the folded structure.

In these circumstances, it is important to run a western blot in non-denaturing conditions, and this will be noted on the datasheet in the applications section. In general, a non-denaturing condition simply means leaving SDS out of the sample and migration buffers and not heating the samples. Rule of thumb: reduce and denature unless the datasheet specifies otherwise. Western blot tools.

Primary antibodies for WB. Loading controls. Secondary antibodies optimized for WB. Western blot buffers. Transfer and staining. Membrane stripping. Troubleshooting tips. General protocol. Fluorescent western blotting. Fluorescent WB protocol. For other video protocols please visit our video protocols library here. For proteins larger than 80 kDa, we recommend that SDS is included at a final concentration of 0.

Add to TBST buffer. Mix well and filter. Failure to filter can lead to spotting, where tiny dark grains will contaminate the blot during color development. T he time and voltage may require optimization. A reducing gel should be used unless non-reducing conditions are recommended on the antibody datasheet. The membrane can be either nitrocellulose or PVDF. Activate PVDF with methanol for 1 min and rinse with transfer buffer before preparing the stack.

The time and voltage of transfer may require some optimization. Transfer of proteins to the membrane can be checked using Ponceau S staining before the blocking step. The purpose of western blotting is to separate proteins on a gel according to the molecular weight.

The proteins are then transferred onto a membrane where they can be detected using antibodies. Heat the samples and 95 degrees C for five to 10 minutes in a sample buffer containing a reducing agent such as beta-mercaptoethanol. This results in linearized proteins with a negative charge proportional to their size.

Place a gel into the electrophoresis tank and add in buffer, ensuring the tops of the wells are covered. Acrylamide percentage of the gel being used depends on the molecular weight of the target protein.

Load a molecular weight market into the first lane then load the samples into adjacent wells. All the samples contain equal amounts of protein. Once all the samples are loaded, add running buffer, place the lid onto the electrophoresis tank. Turn on the power supply and set the voltage recommended by the manufacturer of the gels in the gel tank. You should be able to see bubbles rising through the tank. Run the gel until the dye front has moved sufficiently down the gel. The next stage is to transfer the proteins from the gel onto a membrane.

Membranes are usually made from nitrocellulose or PVDF. Remove the gel from the tank and carefully release it from its plastic case. Cut off the wells and the gel foot and place the gel into transfer buffer. Prepare the transfer stack by sandwiching the membrane and gel between filter paper and sponges. The membrane should be closest to the positive electrode and the gel closest to the negative electrode.

Use a small roller to remove any bubbles between the gel and the membrane. Clamp the transfer case closed and submerge it into a transfer tank containing transfer buffer. Add water to the outer chamber to keep the system cool and put on the lid. Turn on the power supply to begin protein transfer. Time and voltage require optimization, so check the manufacturer's instructions for guidance. Now that the proteins have migrated from the gel onto the nitrocellulose membrane, the protein of interest can be detected with an antibody.

The membrane can be removed from the cassette and the molecular weight marker should now be visible.