Western blot protocol

Western blotting is a widely used technique for detecting specific proteins in complex samples. The western blotting procedure refers to the comprehensive process described in this protocol, encompassing all steps from protein separation to detection. This protocol outlines the complete workflow, from sample preparation to detection, using chemiluminescent or fluorescent methods. It includes detailed steps for lysate preparation, gel electrophoresis, protein transfer to membranes, antibody incubation, and imaging. The method relies on separating proteins by size using SDS-PAGE, transferring them to a membrane, and probing with antibodies specific to the target protein. This guide is suitable for both cell culture and tissue samples and supports various detection systems.

Introduction

Western blotting is a cornerstone technique in molecular biology and biochemistry for identifying and quantifying target proteins. It combines gel electrophoresis with immunodetection, allowing researchers to analyze protein expression, post-translational modifications, and molecular weight. The method involves electrophoretic separation of proteins by size, transferring them to a membrane, and detecting them using specific antibodies. This protocol is designed to help researchers achieve high-quality results with minimal background noise. It includes optimized steps for sample preparation, gel running, transfer, blocking, antibody incubation, and detection. Suitable for a wide range of sample types, this guide supports both chemiluminescent and fluorescent detection systems, making it adaptable to various experimental needs.

Background and principles

Western blotting operates on the principle of protein separation by size through SDS-PAGE, which is a form of polyacrylamide gel electrophoresis used to resolve proteins from a complex mixture. In this process, sodium dodecyl sulfate is used to denature proteins and impart a uniform negative charge, resulting in negatively charged proteins. This denaturation disrupts the native protein structure, ensuring that proteins are separated based solely on molecular weight rather than shape or charge, which is crucial for accurate analysis. The denatured proteins are then loaded onto the gel, and electrophoretic separation occurs as an electric current drives their migration; smaller proteins move faster, allowing size-based resolution. After electrophoresis, proteins are transferred to a nitrocellulose or PVDF membrane using an electric field. Blocking agents prevent non-specific binding, and primary antibodies target specific proteins. Secondary antibodies conjugated to enzymes or fluorophores enable visualization. Detection is achieved via chemiluminescence or fluorescence, producing bands that indicate protein presence and quantity. This method is highly specific and sensitive, making it ideal for protein analysis.

Stage 1 - Sample preparation

Before running a western blot, we must make our protein of interest accessible to the antibodies. This usually involves preparing a lysate containing the proteins from cells or tissues. Lysates can be diluted into several aliquots in a loading buffer and stored frozen at -80°C until ready for use.

Materials required

• Your sample

• Lysis buffer (example RIPA ab156034, non-denaturing ab152163)

• PBS

• Protease inhibitor cocktail (example ab65621)

• Phosphatase inhibitor cocktail (for phosphorylated proteins - example ab201112)

• Concentrated loading buffer

• Dithiothreitol (DTT) (example ab141390)

• A BCA or Bradford assay kit (ab102536, ab102535)

Steps

Prepare a lysis buffer according to the manufacturer’s instructions.

• If not included, add protease inhibitors to the lysis buffer. Include phosphatase inhibitors for phosphorylated proteins.

Isolate your cells and suspend them in lysis buffer.

• You should prepare a suspension with ~ 1 mL of lysis buffer added for every 1x10⁷ cells.

• For suspension cells:

  • Wash cells twice with PBS by spinning down (100–500 g, 5 min, 4°C) and resuspending the pellet.
  • Spin down again (100–500 x g, 5 min, 4°C) and resuspend in ice-cold lysis buffer.

• Adherent cells may require enzymatic or mechanical detachment prior to washing, spinning down (100–500 x g, 5 min, 4°C), and resuspending the pellet in lysis buffer.

Lyse the cell suspension.

• Incubate the cells in lysis buffer for 10 min at 4°C, with rocking.
• Sonicate the suspension to break open cells.

Spin down the suspension to pellet insoluble contents.

• Centrifuge the suspension at 14,000–17,000 x g for 5 min at 4°C.

Keep the supernatant in place in a fresh tube on ice.

• The pellet can be discarded.

Determine the protein concentration of your lysate using a Bradford or BCA assay.

Aliquot the lysate into several tubes.

• Pipette the same volume into each tube.

Dilute the aliquots in loading buffer.

• Ensure the loading buffer contains DTT.
• Add loading buffer to dilute aliquots to a total protein concentration of around 1–2 mg/mL.

Store samples at -80°C until ready for use.

Materials required

• Your sample

• Lysis buffer (example RIPA ab156034, non-denaturing ab152163)

• PBS

• Protease inhibitor cocktail (example ab65621)

• Phosphatase inhibitor cocktail (optional - example ab201112)

• Tubes loaded with glass beads

• Automated homogenizer

• Concentrated loading buffer

• Dithiothreitol (DTT) (example ab141390)

• A BCA or Bradford assay kit (ab102536, ab102535)

Steps

Prepare a lysis buffer according to the manufacturer’s instructions.

• If not included, add protease inhibitors to the lysis buffer. Include phosphatase inhibitors for phosphorylated proteins.

Dissect the tissue with clean tools on ice.

Place dissected tissue and lysis buffer in tubes loaded with glass beads.

• For each 200 mg piece of tissue, add 1,200 µL of lysis buffer.

Lyse the tissue suspension using an automated homogenizer.

• Homogenize the suspension for ~ 3 min at 4°C.
• Incubate for around 5 min at 4°C.

Spin down the suspension to pellet insoluble contents.

• Centrifuge the suspension at 14,000 - 17,000 g for 5 - 10 min at 4°C.

Keep the supernatant in place in a fresh tube on ice.

• This is your lysate.
• The remaining pellet can be discarded.

Determine the protein concentration of lysate using a Bradford or BCA assay.

Aliquot the lysate into several tubes.

• Pipette the same volume into each tube.

Dilute the aliquots in loading buffer.

• Ensure the loading buffer contains DTT.
• Add loading buffer to dilute aliquots to a total protein concentration of around 1–2 mg/mL.

Store samples at -80°C until ready for use.

Stage 2 - Loading and running the gel

The gel is immersed in buffer, the protein samples are loaded, and an electrical current is applied to the gel, which causes proteins to migrate from one end of the gel (negative electrode) to the other (positive electrode). Proteins are separated by size; smaller proteins travel more quickly through the gel, so appear further down.

To confirm the size of each protein in your sample, they are run alongside molecular weight ladders.

Materials required

• SDS-PAGE gel (Tris-Glycine, Bis-Tris or Tris Acetate based gel)

• Molecular weight ladder (example ab116028)

• Gel running apparatus

• SDS

• Running buffer (example Tris-Glycine, MES, MOPS, Tris-Acetate)

Steps

Select an appropriate SDS-PAGE gel for your protein and set up the running apparatus.

• Select or prepare a gel and buffer system based on the protein’s size.
• Place your gel into the running apparatus and fill it with running buffer so that the gel is fully immersed.

Table 1: Recommended gradient gel chemistries for different protein sizes. Our lab use gradient gels, but gels with fixed acrylamide concentration can also be used.

Table 2: Recommended gel chemistries to use for fixed-concentration Tris-Glycine gels. Some optimization will be required if preparing your own gels; a 10 - 15% separating gel is often a good starting point.

Thaw and fully denature your lysates.

• Thaw your lysate, then store it on wet ice.
• Boil each lysate at 100°C for 10 min.

Load an equal quantity of protein from each sample into the gel.

• We recommend:

  • 10−40 µg of protein from a lysate
  • 10−500 ng of purified protein

Run the gel according to the manufacturer's instructions.

• Optimize running times and voltages according to the machine you’re using and the target protein.

Remove the gel from the running apparatus when ready to transfer.

• Use a gel knife to carefully pry open the apparatus and remove the gel.

Stage 3 - Transferring from the gel to the membrane

After performing electrophoresis, proteins are then transferred (or ‘blotted’) onto a membrane, ready for antibody incubation. This membrane can be made of nitrocellulose or PVDF; either material is acceptable.

As with electrophoresis, transfer to the membrane is achieved by applying an electrical charge, which causes the proteins to migrate. The proteins travel away from the gel near the negative electrode and towards the positive electrode, where they bind to the membrane.

Semi-dry transfer requires additional equipment but has a much shorter transfer time with easier setup.

Materials required

• Your SDS-PAGE gel

• Transfer apparatus

• Wash buffer (example TBST)

• Membrane (either nitrocellulose or PVDF – examples: ab133411, ab133412, and ab133413)

• Methanol (if using PVDF)

Steps

Soak the membrane in methanol, if using a PVDF membrane.

Soak the membrane in water, then in transfer buffer for 10 min at 4°C.

Assemble the SDS-PAGE gel and the membrane in the transfer cassette.

• The gel should be closest to the negative electrode.
• The membrane should be closest to the positive electrode.
• Press down on the stack with a small roller to remove any bubbles.

Run the transfer according to the manufacturer's instructions.

Remove the gel and membrane from the transfer apparatus.

Wet transfer is performed in a tank. It doesn't require as much additional equipment as semi-dry transfer, but the transfer time is typically longer.

Materials required

• Your SDS-PAGE gel

• Transfer apparatus

• Transfer buffer (example Tris-Glycine)

• Wash buffer (example TBST)

• Membrane (either nitrocellulose or PVDF – examples: ab133411, ab133412, ab133413)

• Methanol (if using PVDF)

Steps

Soak the membrane in methanol, if using PVDF.

Soak the membrane in water, then in transfer buffer for 10 min at 4°C.

Assemble the SDS-PAGE gel and the membrane in the transfer apparatus.

• The gel should be closest to the negative electrode.
• The membrane should be closest to the positive electrode.
• Press down on the stack with a small roller to remove any bubbles.

Run the transfer according to the manufacturer's instructions.

Remove the gel and membrane from the transfer apparatus.

Stage 4 - Checking the success of transfer (optional)

Before proceeding, you can check the protein has successfully transferred to the membrane.

You can check the success of the transfer using Coomassie staining of the gel.

You can use the pre-stained molecular weight ladder as an initial check to compare the amount of protein on the PAGE gel and the membrane.

For the best quality results, Ponceau S staining is not recommended for fluorescent western blot because it can lead to high background fluorescence, even after extensive washing. There are alternative protein stains that do not fluoresce.

Materials required

• Your membrane

• Your SDS-PAGE gel

• Coomassie stain (example ab119211)

Steps

Observe the colored bands of the pre-stained molecular weight ladder.

• Colored bands should be clearly visible on the membrane.

Stain the SDS-PAGE gel in Coomassie stain.

• Immerse the gel in Coomassie stain and incubate according to the manufacturer’s instructions.
• Wash extensively with water until the background is removed.
• Blue bands indicate proteins remaining on the gel.

Stage 5 - Blocking and antibody incubation

Use the procedures below for antibody incubations. If using loading control antibodies in chemiluminescent western blot, the staining procedure below can be repeated on the same membrane after stripping.

In fluorescent western blot, the membrane can be incubated with multiple sets of antibodies simultaneously according to the following procedure. Loading control antibodies and detection antibodies can be run on the same membrane without the need for stripping.

Materials required

• Blocking buffer (example: 3–5% milk or BSA in TBST, or non-mammalian protein buffer)

• Wash buffer (example TBST)

• Your membrane

• Conjugated primary antibody

Steps

Place the membrane in a container and cover with blocking buffer.

• For fluorescent western blot, incubate the membrane with gentle rocking for 1 h at room temperature.
• For chemiluminescent western blot, incubate the membrane with gentle rocking overnight at 4°C, or for 1 h at room temperature

Dilute the antibody in blocking buffer to the recommended dilution.

Cover the membrane with primary antibody in blocking buffer.

• Incubate with gentle rocking overnight at 4°C, or 1 h at room temperature.

Wash the membrane three times with wash buffer, 5 min each.

Materials required

• Blocking buffer (example: 3–5% milk or BSA in TBST, or non-mammalian protein buffer)

• Wash buffer (example TBST)

• Your membrane

• Primary antibody

• Conjugated secondary antibody

Steps

Place the membrane in a container and cover with blocking buffer.

• For fluorescent western blot, incubate with gentle rocking for 1 h at room temperature.
• For chemiluminescent western blot, incubate the membrane with gentle rocking overnight at 4°C, or 1 h at room temperature.

Dilute the antibody in blocking buffer to the recommended dilution.

• Optimum dilutions will often be suggested on the antibody datasheet.

Cover the membrane with primary antibody in blocking buffer.

• Incubate with gentle rocking overnight at 4°C, or 1 h at room temperature.

Wash the membrane three times with wash buffer, 5 min each.

Cover the membrane with conjugated secondary antibody in blocking buffer.

• Incubate with gentle rocking for 1 h at room temperature.

Wash the membrane three times with wash buffer, 5 min each.

Stage 6 - Detection

Once incubation is complete, you’re now ready to image your western blot.

In chemiluminescent detection, the first step is to incubate the blot in a chemiluminescent substrate solution, which will cause light to be emitted where HRP-conjugated antibodies are present. These bands of light on the blot correspond to antibody binding and should be resolvable as bands on the blot. Chemiluminescent blots have been traditionally imaged using X-ray film-based techniques, but these are largely being replaced by benchtop charge-coupled device (CCD) imagers, which provide much higher quality images.

Materials required

• Your blot stained with conjugated antibodies (eg, Alexa-Fluor^® antibodies)
• 70% Ethanol
• Cotton lint-free cloth
• Silicon mat
• Filter paper (if scanning dry)
• TBST (if scanning wet)
• Imaging system

Alexa Fluor^® is a registered trademark of Life Technologies. Alexa Fluor^® dye conjugates contain(s) technology licensed to Abcam by Life Technologies.

Steps

Clean the imaging scanning bed with 70% ethanol using a cotton lint-free cloth.

Scan membranes in the imaging system.

• To scan membranes wet:

  • Place membranes on scanning bed protein side down.
  • Spray ~ 5 mL of TBST across the scan area.
  • Place a silicon mat on the membranes and use a roller to remove any bubbles.
  • Close lid and scan according to the equipment’s instructions.

• To scan membranes dry.

  • Place membrane between two sheets of filter paper, cover with foil and leave overnight to dry.
  • Place membranes on scanning bed protein side down.
  • Place a silicon mat on the membranes to keep them in place.
  • Close lid and scan according to the equipment’s instructions.

Remove the membranes from the scan bed and clean the imaging scan bed with 70% ethanol using a cotton lint-free cloth.

If available, we recommend the use of charge-coupled devices (CCDs) to image western blots according to the procedure below.

Materials required

• Your blot incubated with HRP-conjugated antibodies (for example - HRP antibodies)
• Chemiluminescent substrate (for example – ECL substrate kits)
• Chemiluminescence imaging system
• Tissue paper

Steps

Prepare the chemiluminescent substrate solution as recommended by the manufacturer.

Incubate the blot in the substrate solution for up to 5 min.

• Follow the manufacturer's advice for more detail.

Remove excess substrate by dabbing the edge of the blot with tissue paper.

Expose your blot using your chemiluminescence imaging system.

• Place the blot on the imaging tray.
• Set up the imaging system to take an image every 30 seconds for up to 20 min.

Although our in-house scientists no longer image using X-ray techniques, the following procedure may be useful to researchers without access to a CCD imaging system. Note that X-ray film requires development in a darkroom.

Materials required

• Your blot incubated with HRP-conjugated antibodies (example - HRP antibodies)
• Chemiluminescent substrate (example – ECL substrate kits)
• X-ray film
• Tissue paper

Steps

Prepare the chemiluminescent substrate solution as recommended by the manufacturer.

Incubate the blot in the substrate solution for up to 5 min.

• Follow the manufacturer's advice for more detail.

Remove excess substrate by dabbing the edge of the blot with tissue paper.

Expose your blot using to X-ray film in a darkroom.

• Expose the blot to film using the following exposure times as a guide in the first instance:

  • 10 s
  • 30 s
  • 1 min
  • 5 min (if no signal apparent at 1 min)
  • 10 min

Stage 7 - Membrane stripping (optional)

Stripping the membrane allows you to remove antibodies from the membrane and restain it with a different set of antibodies. This is helpful if you intend to incubate the membrane with loading control antibodies.

Materials required

• Stripping buffer (for example, ab282569)
• TBST
• PBS
• Your membrane
• ECL detection reagent

Steps

Strip the membrane with stripping buffer.

• Incubate in stripping buffer for around 20 min.

Wash the membrane in PBS for 5 min at room temperature.

Incubate the membrane with a small amount of ECL detection reagent.

• Incubate for around 5 min to check the stripping has been successful.
• If the membrane is clear, proceed to blocking and antibody incubation.

Stage 8 - Data analysis

Proteins can be identified by bands at or near the expected molecular weight, as confirmed by the molecular weight ladder. To rule out non-specific interactions, the same band should be absent in the negative control lane. For example, in the figure above, we see that the negative control lane does not have a band for the target (CD133).

Note that bands can differ from the expected molecular weight for a range of reasons, including:

• Post-translational modifications, such as phosphorylation and glycosylation, increase the size of the protein.
• Splice variants and isoforms may create different-sized proteins produced from the same gene.
• Relative charge: The composition of amino acids can affect how far the protein will travel through the gel.
• Multimers: This is usually prevented in reducing conditions, although strong interactions can result in the appearance of higher bands.

If the bands are at an unexpected molecular weight or difficult to resolve in any other way, please refer to our troubleshooting guide.

Proteins can be identified by bands at or near the expected molecular weight, as confirmed by the molecular weight ladder. To rule out non-specific interactions, the same band should be absent in the negative control lane.

For example, in Figure 4, we see a band for vinculin at 124 kDa in wild-type (Lane 1: wild-type A431 whole cell lysate) and HeLa (Lane 3: HeLa whole cell lysate) samples. However, it is absent in the negative control lanes (Lane 2: vinculin knockout A431 whole cell lysate, and Lane 4: Jurkat whole cell lysate).

Note that bands can differ from the expected molecular weight for a range of reasons, including:

• Post-translational modifications, such as phosphorylation and glycosylation, increase the size of the protein.
• Splice variants and isoforms may create different-sized proteins produced from the same gene.
• Relative charge: The composition of amino acids can affect how far the protein will travel through the gel.
• Multimers: This is usually prevented in reducing conditions, although strong interactions can result in the appearance of higher bands.

If the bands are at an unexpected molecular weight or difficult to resolve in any other way, please refer to our troubleshooting guide.

If you have included loading controls, it’s possible to estimate the relative expression of proteins in your samples. This is done using image analysis software which can measure the brightness of bands relative to their background.

Materials required

• An image of your western blot
• Suitable image analysis software

Steps

Using image analysis software, measure the brightness of the following bands:

• Bands for the protein of interest in each lane (P)
• Loading control bands in each lane (LC)

Measure the brightness of an area of background (B) just below each of the bands measured in step 1.

• Subtract this background from the brightness of the respective band measured in step 1.

Divide the normalized brightness of the protein of interest (P - B1) by the normalized intensity of the loading control (LC – B2).

Relative expression = P – B1 divided by LC – B2

This will give you the relative expression of your protein of interest to your loading control in each lane.

Comparison to other methods

Compared to techniques like enzyme linked immunosorbent assay (ELISA) or mass spectrometry, western blot analysis offers a balance of specificity, sensitivity, and visual confirmation. ELSIA provides highly specific and quantitative data for detecting proteins, antibodies, or antigens in biological samples, but lacks size resolution. Mass spectrometry delivers detailed protein identification but requires complex instrumentation and expertise. Immunohistochemistry allows localization within tissues but is less quantitative. Western blot analysis stands out by combining size-based separation with antibody specificity, enabling detection of post-translational modificationsand isoforms. It is more accessible than mass spectrometry and more informative than ELISA when protein size matters. While not as high-throughput, western blot analysis remains a gold standard for validating protein expression and assessing molecular weight.

Applications

Western blotting is used across biomedical research, diagnostics, and biotechnology. It enables detection of specific target proteins in cell lysates or tissue extracts, making it essential for studying gene expression, signaling pathways, and disease biomarkers. Researchers use it to confirm antibody specificity, validate knockdown or overexpression experiments, and monitor protein modifications like phosphorylation. Immunodetection in western blotting allows for the identification of the target antigen within complex samples. It supports both qualitative and semi-quantitative analysis. In clinical settings, western blotting aids in diagnosing infections and autoimmune disorders. Its adaptability to different sample types and detection systems makes it a versatile tool in proteomics, cell biology, and molecular diagnostics. The protocol supports applications from basic research to translational studies, including the study of protein interactions and binding properties.

Limitations

Despite its strengths, western blotting has limitations. It is time-consuming and requires careful optimization of each step, from sample preparation to detection. Sensitivity can be affected by antibody quality, transfer efficiency, and blocking conditions. Quantification is semi-quantitative at best, and results may vary between experiments. Detection of low-abundance proteins may require signal amplification or enrichment strategies. Membrane handling and imaging can introduce variability. Additionally, the technique is limited to denatured proteins that can be separated by SDS-PAGE. It does not provide spatial information like immunohistochemistry or comprehensive profiling like mass spectrometry. Proper controls and validation are essential for reliable interpretation.

Troubleshooting

Common issues in western blotting include weak signals, high background, and distorted bands. Weak signals may result from low protein concentration, poor antibody binding, or inefficient transfer. Ensure proper lysis, accurate quantification, and optimized antibody dilutions. High background often stems from inadequate blocking or excessive antibody concentration. Use appropriate blocking buffers and wash thoroughly. Distorted or smeared bands can be caused by overloading wells, uneven gel polymerization, or incorrect running conditions. Verify gel composition and loading volumes. If transfer is incomplete, check membrane activation and transfer settings. Consistent sample handling, clean equipment, and validated reagents are key to troubleshooting and improving results.