Western blotting guide

Western blotting is a powerful analytical technique used to detect specific proteins in a given sample. It combines gel electrophoresis for protein separation with antibody-based detection. This technique is crucial for studying protein expression, size, and post-translational modifications.

Introduced by Towbin et al. in 1979, Western blotting has become a standard and essential method for protein analysis. This technique, also known as protein blotting or immunoblotting, utilizes antibodies to detect specific proteins. These proteins are first separated using electrophoresis and then transferred onto a membrane. The precise antibody-antigen interaction allows for the identification of a target protein within a complex mixture, such as a cell or tissue lysate. Western blotting is capable of providing both qualitative and semi-quantitative information about a protein of interest. In this handbook, you will discover guidance on selecting appropriate protein transfer and detection techniques for Western blots. Additionally, it includes troubleshooting advice, instructional videos, selection guides, and buffer recipes to assist in achieving the best possible results.

Sample preparation is key for successful Western blotting. This involves cell lysis to extract proteins, followed by quantification to ensure equal protein loading. Proper preparation avoids degradation and ensures reliable results.

Western blotting involves preparing samples that include lysis buffers, lysates from cell cultures, and tissue lysates, as well as determining protein concentration. The sample preparation consists of three primary steps:

1. Lysis: Cells or tissue samples are lysed using buffers and mechanical agitation to release proteins.
2. Protein Concentration Measurement: This step involves quantifying the protein content.
3. Reducing and Denaturing: Typically, samples are treated with agents to disrupt higher-order structures like dimers and tertiary structures.

• On ice, wash cells in a culture dish with ice-cold PBS.
• Add ice-cold lysis buffer (volume based on cell and dish size), aspirate the PBS, and scrape off cells with a cold plastic scraper. For trypsinized cells, resuspend in lysis buffer in a microcentrifuge tube.
• Agitate for 30 minutes at 4°C and centrifuge. The centrifugation parameters depend on cell type.
• Transfer the supernatant to a new tube on ice and discard the pellet.

• Dissect tissue quickly on ice to prevent protease degradation.
• Snap freeze the tissue in liquid nitrogen and store at -80°C or process immediately.
• Add ice-cold lysis buffer to the tissue in a tube, homogenize, and agitate for 2 hours at 4°C.
• Centrifuge and transfer the supernatant to a new tube on ice. Discard the pellet.

• Measure the protein concentration using Bradford, Lowry, or BCA assays with BSA as a standard.
• Determine the sample concentration, then freeze or prepare for immunoprecipitation or gel loading.

• Denature and reduce samples using a loading buffer with SDS and heat.
• The standard loading buffer is 2X Laemmli buffer, which can be mixed in different strengths.
• Glycerol and bromophenol blue are added to the loading buffer for density and migration tracking, respectively.
• Vortex the sample before and after heating for optimal resolution.

• For epitopes in native structures, run the Western blot under non-denaturing conditions, omitting SDS and not heating the samples.
• Certain antibodies require non-reduced protein; omit reducing agents in these cases.

In this step, proteins are separated based on size through gel electrophoresis. Polyacrylamide gels are commonly used, and the choice of gel percentage depends on the protein size. Proper electrophoresis conditions are critical for resolution and separation.

Electrophoresis refers to the movement of charged molecules through a solvent under the influence of an electric field. This technique is an efficient, quick, and sensitive method for separating proteins and nucleic acids, as any ion or molecule with a charge will migrate in an electric field. Biological molecules, which typically have a net charge at pH levels different from their isoelectric point, move at a rate corresponding to their charge density.The factors influencing a biological molecule’s mobility in an electric field include:

• Field strength.
• The molecule’s net charge.
• Its size and shape.
• Ionic strength.
• Characteristics of the matrix it moves through, such as viscosity and pore size.

The acrylamide matrix in gel electrophoresis can vary, leading to different gel types such as linear vs. gradient gels. Linear gels have a single acrylamide percentage, while gradient gels feature a range of percentages, allowing for the separation of a wider protein range.Gels can also be classified as continuous or discontinuous. A continuous gel is formed from a single acrylamide solution throughout the entire gel cassette, whereas a discontinuous gel uses two different acrylamide solutions: a lower percentage stacking gel and a separating gel. In the Tris-glycine system, proteins are concentrated in the stacking gel for better resolution when they reach the separating gel.Electrophoresis gels come in two sizes: mini and midi. Midi gels are wider, allowing for more or larger wells, facilitating the loading of more samples or larger volumes.Buffer systems in electrophoresis are either continuous, using the same buffer in both the gel and running buffer, or discontinuous, employing different buffers in each. The discontinuous buffer system can include two gel layers with distinct pore sizes and compositions, improving sample concentration and resolution.Electrophoresis conditions also vary, including:

• Denaturing conditions (SDS-PAGE), where proteins are denatured using sodium dodecyl sulfate (SDS).
• Nondenaturing conditions (native PAGE), maintaining native protein structures.
• Reducing conditions, using agents like dithiothreitol (DTT) to unfold denatured proteins into subunits.

After electrophoresis, proteins are transferred from the gel to a membrane, typically PVDF or nitrocellulose. The transfer can be done using either wet or semi-dry methods, ensuring that proteins are immobilized on the membrane for detection.

The process of protein transfer in Western blotting is pivotal for enhancing handling capabilities and ensuring effective target accessibility for macromolecules such as antibodies. This transfer is achieved through the electrophoretic mobility of proteins, enabling their movement from a polyacrylamide gel onto a more stable membrane, typically made of nitrocellulose or polyvinylidene difluoride (PVDF). The procedure involves sandwiching the gel and membrane between electrodes, submerged in a transfer buffer, and applying an electric field to facilitate protein movement onto the membrane surface.

Western blotting employs electrotransfer, specifically by one of three primary techniques: wet (or tank) transfer, semi-dry transfer, and dry transfer. These methods differ mainly in the amount of buffer used and the speed of the transfer.

Wet transfer, the most conventional method, involves immersing the transfer sandwich in a buffer tank and applying either constant current or voltage. This method is particularly effective for lower molecular weight proteins but poses a risk of ‘blow-through’ for proteins under 30 kDa with prolonged transfer times and larger pore-size membranes.

Semi-dry transfer, developed for quicker results, uses limited buffer contained within the transfer sandwich, placed horizontally between plate electrodes. This method emphasizes the importance of ensuring the membrane and filter paper are precisely sized and equilibrated in the transfer buffer. Semi-dry techniques often use higher ionic strength buffers and a high-current power supply to expedite the transfer process.

Lastly, dry transfer eliminates the need for traditional transfer buffers by using a unique gel matrix (transfer stack) incorporating the buffer. This method, characterized by rapid transfer and reduced blot distortion due to the absence of oxygen generation during the process, represents a significant time-saving advancement in Western blotting techniques.

Choosing the right membrane is crucial in Western blotting, and this decision largely depends on the specific needs of the experiment.

Nitrocellulose Membranes: These membranes are highly favored due to their strong protein-binding affinity and versatility with various detection methods. Protein attachment to nitrocellulose is believed to occur through hydrophobic interactions. Optimizing transfer conditions with higher salt concentrations and lower methanol levels can enhance protein immobilization, particularly for higher molecular weight proteins. Key characteristics of nitrocellulose membranes include:

• Superior Quality: Pure, 100% nitrocellulose with a high surface area and consistent quality.
• Range of Choices: Available in 0.2 µm and 0.45 µm pore sizes, suitable for peptides and proteins respectively.
• Enhanced Sensitivity: These membranes facilitate high-affinity protein binding, easy blocking, and minimal background in chemiluminescent Western blotting.

PVDF Membranes: Ideal for Western blotting, amino acid analysis, and protein sequencing (even with minute quantities), PVDF membranes are highly hydrophobic and require pre-wetting with methanol or ethanol. They exhibit strong protein binding affinity, primarily through dipole and hydrophobic interactions, and are superior in retaining adsorbed proteins compared to other materials. PVDF’s resilience against brittleness allows for multiple reprobings without losing sensitivity or increasing background noise. Key features include:

• Robust Quality: PVDF membranes designed specifically for protein transfer and Western blot applications, exhibiting resistance to discoloration.
• Durability: Compatible with a variety of organic solvents, acids, and mild bases, and less prone to tearing or brittleness than nitrocellulose.
• Broad Selection: Offered in 0.2 µm and 0.45 µm pore sizes, in various formats such as pre-cut sheets, rolls, and device-specific formats.
• Versatility: Suitable for different detection methods including chemiluminescent, chromogenic, and fluorescent.

For Western blotting and other applications, the 0.2 µm PVDF membrane is reliable, while the 0.45 µm version is particularly suited for high-sensitivity and low-background immunoblotting.

Blotting Paper: In wet and semi-dry transfer methods, blotting (or filter) paper is a vital component. It is soaked in transfer buffer to help proteins migrate from the gel to the membrane. The quality of the blotting paper is important to avoid background issues during detection. The paper’s thickness plays a significant role, especially in semi-dry transfers, where thicker paper can hold more buffer and facilitate better transfer. It’s important to adhere to the specific thickness recommendations for filter paper provided by the transfer device manufacturer. Dry electrotransfer methods do not utilize blotting paper.

Blocking non-specific sites on the membrane is essential to prevent non-specific binding. This is followed by incubation with primary antibodies specific to the target protein, and then with secondary antibodies for detection.

In Western blotting, membranes are highly attracted to proteins, necessitating the use of blocking buffers to prevent non-specific antibody binding after transfer. Common blocking agents include milk and bovine serum albumin (BSA). Milk is a versatile blocker, effective for general applications. However, for detecting phosphoproteins, BSA is preferred as it avoids the phosphoproteins often present in milk, which can cause false positive signals.

Balancing the concentration of blocking agent is crucial. Insufficient amounts lead to high background noise and a low signal-to-noise ratio, while excess can mask antibody-target interactions or inhibit marker enzymes, reducing signal clarity. The optimal blocking buffer often depends on the specific system used in Western blotting. For example, with alkaline phosphatase conjugates, Tris-buffered saline (TBS) is recommended over phosphate-buffered saline (PBS), which interferes with the enzyme activity. Conversely, milk should be avoided in avidin-biotin systems. Experimentation with various blocking buffers, including milk and BSA, is key to achieving the best results in your specific setup.

Primary antibodies. Western blot analysis largely relies on target-specific probes, detectable through chemical tags or labels, to identify specific proteins in biological samples. Antibodies, due to their high affinity for particular proteins, are the most common type of probe. They can locate and detect specific proteins in complex samples, such as cell or tissue lysates. However, antibodies, being proteins themselves, require tagging for visualization (primary conjugates) or need to be used with a tagged secondary antibody for detection in assays.

Various chemical labels or tags, like enzymes and fluorophores, can be attached to antibodies for detection and measurement. Enzymatic tags, such as alkaline phosphatase (AP) and horseradish peroxidase (HRP), are commonly used in chemiluminescence methods for western blot detection. Meanwhile, fluorescence-based detection methods are gaining traction.

The selection of a primary antibody for western blotting depends on the target protein and available antibodies. Numerous commercial primary antibodies exist, and custom antibodies can also be developed. Both polyclonal and monoclonal antibodies are effective for western blotting. Polyclonal antibodies are less costly and quicker to produce, often displaying high antigen affinity. Monoclonal antibodies are preferred for their specificity, purity, and consistent low background. Crude preparations like serum or ascites fluid are sometimes used, but their impurities may increase background noise. Recombinant antibodies, produced through known DNA sequences of the antibody’s variable region, offer consistency and reproducible performance. Affinity purification using the immobilized target molecule can further enhance antibody specificity.

It’s crucial to verify the specificity of primary antibodies, as non-specific antibodies can bind to unintended proteins, resulting in multiple bands on a western blot and complicating data interpretation.

Secondary antibodies. During the secondary antibody incubation step in Western blotting, the secondary antibody binds to a primary antibody already attached to the target protein. Most primary antibodies come from a limited range of host animals and belong to the IgG class, simplifying the selection of suitable, labeled secondary antibodies for various applications. Secondary antibodies can be polyclonal, monoclonal, or recombinant, and are available in different forms like lyophilized or in solution. Typically, this incubation is done for 1 hour at room temperature, although longer periods can be used if necessary.

The dilution range for secondary antibodies varies, from 1:500 for fluorescent detection to 1:500,000 for chemiluminescent detection of abundant targets, depending on factors like target protein abundance and the choice of primary antibody and chemiluminescent substrate. Enzyme-conjugated secondary antibodies often require higher dilutions due to the enzymatic amplification of the signal.

Secondary antibodies are produced by immunizing a host animal with antibodies from another species. For instance, anti-mouse antibodies are generated by injecting mouse antibodies into a non-mouse animal like goats, donkeys, or rabbits. These antibodies typically target a wide range of immunoglobulins from the source species. To achieve specificity and reduce cross-reactivity, secondary antibodies undergo affinity purification and cross-adsorption.

Enzymes like alkaline phosphatase (AP) and horseradish peroxidase (HRP) are commonly used labels for secondary antibodies in Western blots. AP catalyzes reactions that produce color or light, while HRP oxidizes substrates in the presence of hydrogen peroxide to produce a similar effect. HRP is favored for its high turnover rate, stability, and cost-effectiveness.

Fluorescent labels are becoming more popular in Western blotting. These require imaging equipment to capture the fluorescent signal and can be less sensitive than enzymatic methods, but they allow for multiplexing and reduce chemical waste.

Biotin-binding proteins like avidin, streptavidin, or NeutrAvidin™ Protein, used in conjunction with biotin-tagged probes, offer another method for detection. These proteins have a high affinity for biotin and can be labeled with enzymes or fluorophores for various assay systems. Each has its unique properties and limitations, like avidin’s potential for nonspecific lectin binding and streptavidin’s bacterial recognition sequence, which might cause background binding in certain samples. NeutrAvidin™ Protein, a modified form of avidin, avoids these issues.

Detection of the target protein can be achieved through chemiluminescent, colorimetric, or fluorescent methods. The choice depends on the required sensitivity and the available imaging equipment.

Chemiluminescent detection is a preferred method in Western blotting for its high sensitivity and compatibility with film or digital imaging systems. It offers several benefits:

1. Blots can be reprobed for better detection or to identify a second protein.
2. It allows detection and quantification of a wide range of protein concentrations.
3. Provides superior sensitivity compared to other methods.

Chemiluminescent substrates differ from others as the light emitted is transient, only occurring during the enzyme-substrate reaction. This contrasts with chromogenic substrates that leave a stable, colored product. In chemiluminescent blots, the substrate is the limiting factor, and as it depletes, light production wanes. A well-calibrated process ensures stable light emission for hours, facilitating consistent protein detection.

HRP chemiluminescent substrates usually involve a two-component system, mixing stable peroxide and enhanced luminol solutions. This reaction emits light at 425 nm, captured by X-ray film or imaging instruments. Western blot imaging instruments provide benefits like quantitative analysis and adjustable exposure times, offering more sensitivity and resolution than film. The choice of HRP substrate depends on factors like protein abundance and available equipment.

AP chemiluminescent substrates are alternatives to HRP systems. AP reactions remain linear over time, and longer reactions can enhance sensitivity. AP is not affected by antibacterial agents and operates best at pH 9.0–9.6. AP substrates like CDP-Star offer high sensitivity and are compatible with X-ray film and imaging systems.

Chromogenic detection uses substrates that transform into insoluble, colored products upon contact with enzymes. This simple, cost-effective method is ideal for detecting high-abundance proteins. However, chromogenic substrates have lower sensitivity and can produce background signals over prolonged development times. Unlike chemiluminescent assays, chromogenic blots can’t be easily reprobed.

Chromogenic HRP substrates require peroxide for color development. Commercially available substrates often come with stabilized peroxide, offering consistent results and longer shelf life. TMB and DAB are common substrates, producing blue and brown precipitates, respectively.

AP chromogenic substrates, like Nitro blue tetrazolium (NBT) and BCIP, produce stable, intensely colored precipitates. NBT/BCIP combinations yield a black-purple precipitate with high sensitivity and sharp band resolution.

Fluorescent Western blot detection differs from enzymatic methods as it relies on light emission from excited fluorophores. It offers more stable signals over time, increasing in popularity due to time savings, reduced chemical waste, and advancements in imaging technology. Fluorescent detection allows multiplexing, enabling simultaneous assay of multiple targets on the same blot without reprobing. Despite not matching the sensitivity of chemiluminescent methods, fluorescent detection is increasingly favored in many labs.

The final step involves visualizing the blots using appropriate imaging systems. The analysis includes quantifying band intensity and comparing protein levels across samples.

Chemiluminescence and fluorescence are key methods for capturing data in Western blot detection. Chemiluminescence, driven by HRP or AP reactions, produces light that is captured using X-ray film or Western blot imaging instruments. Capturing chemiluminescent signals with X-ray film requires multiple exposures to balance signal and background, aiming for a clear signal with low background. This process is tricky as it can’t be monitored in real-time, leading to potential underexposure or overexposure, which complicates data analysis. While X-ray film offers qualitative and semi-quantitative data, modern Western blot imaging instruments provide higher sensitivity and a broader dynamic range, allowing for the detection of subtle sample differences. These instruments capture data digitally, eliminating the need for darkroom film processing and subsequent scanning for analysis. They also often include auto-exposure algorithms for optimal signal-to-noise ratios.

Despite the low cost of X-ray film, the expenses related to maintaining a darkroom and development equipment should be considered. In contrast, Western blot imaging instruments, though more costly initially, are versatile, supporting multiple applications including chemiluminescent Western blot signals. This versatility adds value to the investment.

Fluorescence Western blot imaging systems, offered by several manufacturers, capture fluorescence signals using either filter-based or laser-based technologies. Most of these systems use CCD cameras, but photoavalanche diode–based scanning systems are also available. These systems save images digitally and support a growing range of fluorescently labeled antibodies, offering various excitation and emission spectra options.

Fluorophore-based imagers equipped with appropriate filters or lasers enable multiplex analysis, where multiple targets are detected and distinguished on the same blot. This is a significant advantage over chemiluminescence, which requires multiple rounds of stripping and reprobing to visualize each protein of interest. Multiplexing avoids the risk of protein loss during stripping and reprobing, enhancing research efficiency. For instance, it allows simultaneous visualization of a protein of interest with a loading control, differentiation of proteins with similar molecular weights, and evaluation of complex biological pathways. Multiplex experiments are classified based on the number of proteins probed, such as 2-plex for two proteins, 3-plex for three, and so on.

A common protein normalization method involves using housekeeping proteins like α-tubulin, β-actin, or GAPDH as loading controls. However, this approach has limitations, such as variable expression under different experimental conditions and potential signal saturation in Western blots. Optimizing this method requires confirming the linearity of the housekeeping protein signal across a wide concentration range and ensuring it remains unaffected by experimental conditions.

When normalizing posttranslationally modified proteins comparison to their total protein using a pan antibody is often employed. This method, however, faces challenges like ensuring the pan antibody recognizes all modified and unmodified versions of the target and maintaining constant expression of the total protein target across conditions. Additionally, the linear dynamic range for this method is often limited.

Total Protein Normalization (TPN) offers a more reliable alternative. It avoids the variability of housekeeping proteins and the extensive optimization and costs associated with immunodetection reagents. TPN involves labeling or staining the total protein on a blot, comparing and normalizing the relative amounts loaded in each lane to a reference lane. Reversible dyes like Ponceau S are used for TPN, but these require imaging post-staining and destaining before immunoblotting, adding to the time involved.

Another method for TPN involves specialized gels and imagers, which may differ from current optimized systems for protein separation. This method requires imaging the total protein on the blot before immunoblotting, as chemiluminescent signals from target proteins can interfere with the total protein signal.

Common issues in Western blotting include poor transfer, non-specific binding, or weak signal. Troubleshooting often involves optimizing gel electrophoresis, transfer conditions, and antibody concentrations.

Safety in handling samples and reagents is crucial. Best practices include careful preparation, proper documentation, and consistent methodology to ensure reproducibility and accuracy.

Safety in Western blotting is paramount, as the process involves handling various chemicals, biological samples, and equipment that pose potential hazards. To ensure a safe working environment, it is crucial to wear appropriate personal protective equipment (PPE) like gloves, lab coats, and safety glasses. Hazardous chemicals, such as methanol and acrylamide, should be handled with care, and all work should be conducted in a well-ventilated area or under a fume hood to avoid inhalation of harmful vapors. Biological samples, particularly those of human or animal origin, must be treated as potentially infectious and handled following biosafety guidelines. Electrical safety is also important, as electrophoresis and blotting involve high voltage. Equipment should be regularly inspected for any damage, and power sources must be turned off before assembling or disassembling apparatus. Lastly, proper waste disposal protocols must be followed, especially for disposing of hazardous materials, to maintain laboratory safety and environmental compliance.