Here we provide essential tips for the successful transfer and staining of proteins in western blot, including visualization of proteins and development methods for visualization.

Visualization of proteins in western blot

After separating proteins through gel electrophoresis, the next step is to verify if the proteins have migrated uniformly and transfer them onto a specialized membrane. The protein visualization stage is crucial for accessing the quality of protein migration, ensuring uniform and even distribution. Two commonly used methods for protein visualization are copper stain and Coomassie stain.

If we intend to transfer the separated proteins to a membrane, later on, copper stain is recommended. On the other hand, the Coomassie stain is irreversible and fixes the proteins in the gel, making it less suitable for subsequent protein transfer. However, if our main objective is to observe the results of the SDS-PAGE separation without protein transfer, we can employ the Coomassie stain. Alternatively, we can apply the Coomassie stain to gels post-transfer to check for transfer efficiency or use the PVFD membranes for Coomassie staining as they can be destained.

Using Coomassie stain on a PVDF membrane offers several benefits. Firstly, it provides a comprehensive overview of the protein loading on the gel, serving as a potential loading control in addition to any housekeeping proteins that need to be detected. Secondly, after scanning, the PVDF membrane can be effortlessly destained using methanol and automatically activated for further staining.

Using the appropriate staining method and membrane, protein visualization in western blot can effectively evaluate protein migration and loading, ensuring reliable results in downstream analysis.

Coomassie stain

As soon as the power is turned off, the separated protein bands will naturally begin to diffuse since they are freely soluble in an aqueous solution. The crucial step is to prevent protein diffusion by treating the gel with 40% distilled water, 10% acetic acid, and 50% methanol solution. This treatment causes almost all proteins to precipitate, ie, become insoluble.

To visualize the fixed proteins, place the gel in the same solution of water/acetic acid/methanol, but this time add 0.25% Coomassie Brilliant Blue R-250 by weight. Incubate the gel for 4 hours to overnight at room temperature on a shaker. Afterward, transfer the gel (remember to save the dye mixture as it can be reused many times) to a mix of 67.5% distilled water, 7.5% acetic acid, and 25% methanol. Place the gel on a shaker and replace the rinse mixture with a fresh solution until the excess dye has been removed.

The Coomassie stain will not bind to the acrylamide in the gel and will wash out, leaving a clear gel. However, the stain strongly binds to the gel's proteins, resulting in a deep blue color.

Copper stain

Briefly rinse freshly-electrophoresed gels in distilled water for a maximum of 30 seconds and then transfer them to a solution of 0.3 M CuCl2 for 5–15 min. After that, wash the gels briefly in de-ionized water and view them against a dark-field background.

Proteins will appear as clear zones against a translucent blue background. To completely destain the gels, perform repeated washes in a solution of 0.1–0.25 M Tris/0.25 M EDTA pH 8.0. Then, move the gel to a dish of transfer buffer before proceeding with transfer according to the transfer apparatus manufacturer's instructions.

Protein transfer

Protein transfer is the process of transferring proteins from a gel onto a membrane. Manufacturers of transfer apparatus typically provide detailed instructions for the transfer process on their websites. These details vary depending on the system, but the principle remains the same in each case.

Protein transfer is achieved by applying an electrical field, similar to gel electrophoresis, which induces proteins with an electrical charge to migrate from the gel onto a sturdy support, such as a membrane. This process, known as blotting, has evolved from early methods relying on protein diffusion to the now standard practice of blotting in an electrical field, which yields more reliable results.

Transfer can be done using a wet or semi-dry system. The wet transfer is less prone to failure due to membrane drying and is especially recommended for transferring large proteins. For both kinds of transfer, the membrane is placed next to the gel. The two are sandwiched between absorbent materials, and the sandwich is clamped between solid supports to maintain tight contact between the gel and membrane.

Wet protein transfer

During wet protein transfer, the gel and membrane are sandwiched between sponge and paper, arranged in the following order: sponge > paper > gel > membrane > paper > sponge. This sandwich is tightly clamped to ensure no air bubbles are trapped or formed between the gel and membrane. The sandwich is then submerged in the transfer buffer, to which an electrical field is applied. The negatively-charged proteins travel toward the positively-charged electrode but are bound by the membrane, preventing them from further migration.

A standard buffer for wet transfer is the same as the 1x Tris-glycine buffer used as the gel running buffer, but without SDS and with the addition of methanol to a final concentration of 20%. However, for proteins larger than 100 kDa, including SDS at a final concentration of 0.1% is recommended.

Semi-dry protein transfer

In a semi-dry transfer, a sandwich consisting of paper > gel > membrane > paper wetted in transfer buffer is placed directly between positive and negative electrodes (cathode and anode, respectively). Similar to the wet transfer, it is crucial that the membrane is closest to the positive electrode and the gel closest to the negative electrode.

The composition of the transfer buffer for semi-dry transfer may differ from that of wet transfer, so we advise you to consult the apparatus manufacturer's protocol. A standard recipe for semi-dry transfer buffer includes 48 mM Tris, 39 mM glycine, 0.04% SDS, and 20% methanol.

There are two commonly used membrane types: nitrocellulose (eg, 0.45 µm or 0.22 µm membranes) and PVDF (positively charged nylon, eg, low fluorescent western membrane). Both types work effectively, and the choice depends on personal preference. An advantage of the PVDF membrane is that it allows for easy staining and destaining with Coomassie.

Note that PVDF membranes require careful pre-treatment: cut the membrane to the appropriate size and soak it in methanol for 1–2 min. Then, incubate the membrane in an ice-cold transfer buffer for 5 min. Failure to equilibrate the membrane in an ice-cold transfer buffer can lead to shrinking during transfer and a distorted transfer pattern.

Transfer of large and small proteins

Several factors can affect transfer efficiency, like the balance of SDS and methanol in the transfer buffer, protein size, and gel percentage. The following modifications will encourage efficient transfer for different protein types.

Transfer tips for large proteins (>100 kD)

• For large proteins, transfer out of the gel may be very slow, as they run slowly within the gel during separation. If blotting a large protein, run your samples in a low-concentration gel, 8% or less. These will be very fragile, so handle them carefully.
• Large proteins will tend to precipitate in the gel, hindering transfer. Adding SDS to a final concentration of 0.1% in the transfer buffer will discourage this. Methanol tends to remove SDS from proteins, so reducing the methanol percentage to 10% or less will also guard against precipitation.
• Lowering the methanol percentage in the transfer buffer also promotes gel swelling, allowing large proteins to transfer more easily.
• Methanol is only necessary if using nitrocellulose. If using PVDF, methanol can be removed from the transfer buffer altogether and is only needed to activate the PVDF before assembling the gel/membrane sandwich.
• Choose wet transfer overnight at 4°C instead of semi-dry transfer.

Transfer tips for small proteins (100 kD)

• SDS hinders all proteins from binding to membranes, but small proteins are more affected by this than large ones. If your protein of interest is small, omit SDS from the transfer buffer.
• Keep the methanol concentration at 20%.

General transfer tips:

• Avoid touching the membrane with your fingers; use tweezers instead and only touch the edge of the membrane. Oils and proteins on fingers will block efficient transfer and create dirty blots.
• After sandwiching the gel and membrane between paper, we can eliminate air bubbles between the gel and membrane by rolling them out with a roller, pipette, or 15 mL tube. Also, we could assemble the sandwich in a dish of transfer buffer to prevent the formation of bubbles in the first place.
• Make sure the paper and membrane are cut to the same size as the gel. Large overhangs may prevent a current from passing through the membrane in semi-dry transfers.
• Chicken antibodies tend to bind PVDF and other nylon-based membranes, leading to high background. In this case, switching to a nitrocellulose membrane should help reduce background staining.

The following reference discusses a gel and buffer system that allows the transfer of proteins as large as 500 kD: Bolt MW and Mahoney PA (1997). High-efficiency blotting of proteins of diverse sizes following sodium dodecyl sulfate-polyacrylamide gel electrophoresis. Anal Biochem​, 247, 185–92.

Visualization of proteins in membrane with Ponceau Red

Visualizing the proteins using Ponceau Red allows us to check for transfer success. We can implement the following protocol:

1. Wash the membrane in TBST.
2. Dilute the stock of Ponceau Red (example: ab270042 or ab146313) 1:100. The stock is made of 2% Ponceau S in 30% trichloroacetic acid and 30% sulfosalicylic acid.
3. Incubate on an agitator for 5 min, then wash extensively in water until the water is clear and the protein bands are well-defined.
4. The membrane may be destained completely by repeated washing in TBST or water. When using a PVDF membrane, re-activate it with methanol, then wash it in TBST.

Check out the details recipes for TBS 10x and TBST in Buffers and stock solutions for western blot.

TBS 10x

For 1 L;

24.23 g Trizma HCl

80.06 g NaCl

Dissolve in 800 mL distilled water

pH to 7.6 with HCl

Top up to 1 L

TBST
For 1 L;

100 mL TBS 10x

900 mL distilled water

1 mL Tween 20

Tween 20 is very viscous and will stick to the tip of your measuring pipettes. Be sure you add the right amount of the detergent to the Tris buffer. A 10% solution is easier to dispense than undiluted Tween 20.

Blocking the membrane

Blocking the membrane prevents non-specific background binding of the primary and/or secondary antibodies to the membrane, which has a high capacity for binding proteins and, therefore, antibodies.

Two blocking solutions are traditionally used: non-fat milk or BSA (Cohn fraction V). Milk is cheaper but not recommended for studies of phospho-proteins because it contains a phospho-protein casein. Casein will cause high background as the phospho-specific antibodies will detect it.

Some antibodies give a stronger signal on membranes blocked with BSA versus milk for unknown reasons. Check the application notes on the datasheet for specific instructions on how to block the membrane.

To prepare a 5% milk or BSA solution, weigh 5 g per 100 mL TBS with Tween 20 (TBST) buffer. Mix well and filter. Failure to filter can lead to spotting where tiny dark grains will contaminate the blot during development.

Incubate for 1 hr at 4°C under agitation. Rinse for 5 s in TBST after the incubation.

Below are some of abcam's recommended blocking solutions:

• Protein Block (ab64226)
• 10X Blocking Buffer (ab126587)
• Protein Block (ab156024)

Incubation with primary antibody

Incubation buffer

Dilute the antibody in TBST at the dilution suggested in the antibody datasheet. If the datasheet does not specify a dilution, try a range of dilutions (1 : 100 - 1 : 3000 ) and optimize the dilution according to the obtained results. Keep in mind that too much antibody will result in non-specific bands. Certain antibodies may require a different buffer, such as PBS. Regardless of the buffer type selected for the primary antibody, we recommend using the same buffer for the secondary antibody solution.

Some laboratories prefer incubating primary antibodies in a blocking buffer, while others use TBST without a blocking agent. The results vary from antibody to antibody, and you may find it makes a difference to use a non-blocking agent in the antibody buffer or the same agent as the blocking buffer.

If the high background is not an issue, some antibodies produce a much stronger signal if diluted in a buffer with low concentrations (0.5–0.25%) of milk or BSA or even without these additives.

Incubation time

The incubation time can vary between a few hours to overnight (rarely more than 18 h) and depends on the antibody's binding affinity for the target protein and the protein's abundance. We recommend trying a more diluted antibody and a prolonged incubation time to ensure specific binding.

Incubation temperature

​The incubation should be carried out at a cold temperature. If incubating in a blocking buffer overnight, do it at 4°C to avoid contamination and the following protein destruction (especially for phospho groups).

Agitating the antibody solution is recommended to enable adequate homogenous covering of the membrane and prevent uneven binding. During the incubation, ensure you add an adequate solution volume to accommodate your container size. The container itself should be suitable for the size of your membrane, allowing some space. Additionally, covering the container with a lid can help prevent evaporation.

Incubation with secondary antibody

Wash the membrane several times in TBST while agitating, 5 min or more per wash, to remove residual primary antibody. You might also use a pipette to remove residual liquid from the corner of your container.

Incubation buffer and dilution

Dilute the antibody in TBST at the dilution suggested in the antibody datasheet. If the datasheet does not have a recommended dilution, try a range of dilutions (1 : 1,000 – 1 : 2,0000) and optimize the dilution according to the results. Remember that excess of antibody will result in non-specific bands.

To reduce the background, you may incubate the secondary antibody in the blocking buffer. But a background reduction may come at the cost of a weaker specific signal, presumably because the blocking protein hinders the binding of the antibody to the target protein.

Incubation time and temperature

1–2 h at room temperature with agitation.

Which conjugate and target?

We recommend using horseradish peroxidase (HRP)-conjugated secondary antibodies due to their high sensitivity. In contrast, alkaline phosphatase (AP)-conjugated secondary antibodies are less sensitive and not recommended.

It is crucial to select the appropriate type of secondary antibody that matches the type of primary antibody. Various secondary antibody types are available – H&L chain, Fc, heavy chain, mu chain, F(ab´)2, alpha chain, and light chain – which may exhibit different behaviors.

Development methods

Development methods in western blot include using detection kits, X-ray films, and digital images.

Detection kits

For HRP-conjugated antibodies, enhanced chemiluminescence (ECL) kits are traditionally used as substrates. We offer ECL substrate kits with varying detection limits, such as high-sensitivity kits to detect 23 pg–187 ng of protein per band (eg, ab133406) or very high-sensitivity kits to detect 4.6 pg–4.7 ng of protein per band (eg, ab133408).

X-ray films

Many labs work with easy-to-use automated X-ray film developers. Remember that an over-exposed film is unsuitable for analysis as it makes it impossible to determine the relative amount of protein. Over-exposed films show completely black bands with no contrast and/or numerous non-specific bands.

In some cases, multiple exposures might be necessary to identify the optimal exposure time. Weak signals may require longer exposure to the film, while strong signals develop quickly. If signals vary significantly in intensity, exposing them to separate films may be necessary to ensure optimal visualization.

Transferring the marker band carefully and accurately is crucial to ensure reliable data.

Digital images

The new generation of film developers are units with a camera inside an enclosure, removing the need for a darkroom. The camera detects the chemiluminescence emanating from the membrane, transforming the signal into a digital image for rapid analysis with software provided by the detection machine.

Compared to traditional X-ray films, digital imaging offer a significant advantage – the ability to perform preliminary exposure testing without wasting films. Before capturing the image, we can assess signal intensities, adjust the exposure time and select a suitable camera sensitivity. Moreover, we can monitor the signal capture process by using a shorter exposure time and stopping when the desired signal level is reached. Also, the software can sum up the signals from the previous images, aiding in the analysis.

During detection, it is crucial to stop when the signal intensity has reached its maximum. If the entire ECL solution has been used up, the signal might appear inverse, resulting in a white dot in the middle of the band. Many detection systems will automatically stop detection when a signal has reached its maximum on the blot. However, we advise caution as such a strong signal might lead to underexposure of other bands on the same blot, which would have required further exposure.

Various machines are now commercially available, including systems that do not use HRP-conjugated antibodies (ie, chemiluminescence). For example, STORM Analyzers detect fluorescence from fluorochrome-conjugated secondary antibodies, while the Odyssey Infrared Imaging System detects infrared fluorescence.

Return to western blot protocols