Western blotting, also referred to as immunoblotting, was introduced by Towbin et al. in 1976 and is defined as a technique that utilizes electrophoresis to separate and identify specific proteins using antibodies. The first step is to separate proteins, based on molecular weight, on a poly-acrylamide gel. The proteins are then transferred from the gel onto a nitrocellulose or PVDF membrane using an electric current. Once on the membrane, the proteins can be identified using target-specific antibodies, followed by visualization using secondary antibodies and detection reagents. Often the secondary antibody is complexed with an enzyme, which will produce a detectable signal when combined with an appropriate substrate. There are several ways to detect the signal depending on whether it is chemiluminiscent, fluorescent, or chromogenic. Whatever the system used, the intensity of the signal should correlate with the abundance of the antigen on the membrane. This assay can produce qualitative and semi-quantitative data about the target protein.
A step-by-step protocol, with recommendations on reagents and antibodies to perform your western blot, is also outlined on our Western Blotting Protocol page.
Sample preparation for western blot, or any other downstream application, includes extraction of protein from cell or tissue sample using lysis buffers, and diluting the sample in a high density buffer, which will keep the protein from diffusing away when loading the gel. This buffer also includes a dye that will allow tracking the progress of samples during electrophoresis. Several factors must be taken into account while preparing protein samples, such as the source of the protein, protein localization within the cell, amount of protein required, and the proposed downstream applications. These factors are important when considering the type of lysis buffer to be used.
Lysis buffers differ in their ability to solubilize proteins, with those containing sodium dodecyl sulfate (SDS) and other ionic detergents considered to be the harshest and therefore most likely to give the highest yield. There are 4 main considerations when choosing a lysis buffer:
|Recommended Buffer||Protein Location|
|NP-40 or RIPA buffer||Tris HCl|
|NP-40 or RIPA buffer||Subcellular fractionation protocol|
|Whole cell extract||Cytoplasmic|
|Membrane bound||Nuclear and Mitochondrial|
Table 1: buffers based on protein localization
In order to protect the protein samples from being degraded by proteases and/or phosphatases, lysis buffers should always contain protease and phosphatase inhibitors (for phosphoproteins). These inhibitors should be freshly added to the samples. Ready-to-use cocktails of inhibitors are available from Cell Signaling Technology Phosphatase Inhibitor Cocktail (100X) #5870 , Protease/Phosphatase Inhibitor Cocktail (100X) #5872 , Protease Cocktail #5871.
Cultured mammalian cells or soft tissues can be lysed either by using detergent-free buffer coupled with mechanical shearing, or by using detergent-containing buffer in case of membrane or subcellular proteins. Tissue samples, in contrast to cells, require extensive manipulation to break up tissue architecture and solubilize proteins. Often a Dounce homogenizer or mechanical shearing is required to homogenize tissues. For more details on the lysis protocol, please check Western Blotting Application Guide by Cell Signaling Technology.
This procedure is used to separate nuclear, membrane, and cytoplasmic cell fractions using differential centrifugation. Nuclear extraction is used for studying proteins that are bound to the DNA, such as transcription factors or histones. This procedure requires variation to the standard cell lysis protocol to ensure that the cytoplasmic and nuclear compartments of the cell remain separate.
NOTE: The Cell Fractionation Kit #9038 by Cell Signaling Technology makes it simple to extract subcellular proteins. It comes with a straightforward and easy-to-follow protocol outlined for fractionation into cytoplasmic, membrane/organelle, and nuclear/cytoskeletal fractions.
For the study of nuclear and chromatin-associated proteins, an enhanced signal is observed in sonicated extracts compared to nonsonicated extracts (Figure 2). Sonication disrupts membranes and chromatin to a greater extent than lysis buffer alone and ensures complete shearing of chromosomal DNA. Different brands of sonicators perform differently and should be optimized per manufacturer recommendations. If using a probe tip sonicator, we recommend allowing 10 seconds between pulses and keeping samples on ice while sonicating. Bath sonicators are attached to chillers that can be set to desired temperatures, thus eliminating the need for ice. In instances where a sonicator is unavailable, samples can be passed through a fine gauge needle, which will break membranes and shear DNA.
After lysis of cells, it is important to determine the total protein concentration of the sample. Accurate quantitation of the sample will allow you to load the proper amount of protein in each lane. Proper quantitation is also critical when performing semi-quantitative or quantitative westerns.
There are several different methods for quantifying protein concentration in samples. More commonly used assays rely on spectrophotometric determination of protein concentration, such as a Bradford assay, a Lowry assay, or a bicinchoninic acid (BCA) assay. Bovine serum albumin (BSA) is a frequently used protein standard. BCA Protein Assay Kit #7780 is commercially available from Cell Signaling Technology. Once you have determined the concentration of each sample, you can freeze them at -20°C or -80°C for later use or load the samples onto a gel.
Now that the protein samples are ready, it is time to load the samples and run the gel.
It is important to denature the protein with a loading buffer, such as the Laemmli buffer, to be certain that the epitope is accessible. The Laemmli buffer contains anionic detergent sodium dodecyl sulfate (SDS), which coats the peptides with a negative charge. This ensures that the proteins migrate by molecular weight, rather than the intrinsic charge of the polypeptides. Addition of β-mercaptoethanol or DTT to the buffer further reduces disulphide bridges, which is necessary for separation by size. Once the proteins are in the loading buffer, the samples should be boiled at 95°C for 5 min followed by a brief vortex to ensure homogeneity of the sample.
If the antibody in question only recognizes epitopes in the native or non-denatured proteins, SDS needs to be eliminated in the loading and running buffers and the sample should not be heated. Certain antibodies only recognize protein in its non-reduced form (particularly on cysteine residues) such that reducing agents β-mercaptoethanol or DTT must be left out of the loading and running buffers.
The main component of PAGE is the acrylamide gel. Acrylamide, when mixed with bisacrylamide, forms a crosslinked polymer network upon addition of the polymerizing agent, ammonium persulfate (APS). TEMED (N,N,N,N'-tetramethylenediamine) catalyzes the polymerization reaction by promoting the production of free radicals by APS. This mixture creates a sieve-like network that the proteins can then travel through. Gels are available in several different compositions, such as Tris Glycine, Bis-Tris, Tris-Acetate, Tris-Tricine, etc., each of which have different characteristics. For example, Tris-glycine is a general-use gel, suitable for the majority of applications. Bis-Tris HCI gels on the other hand are more acidic, thus offering enhanced stability and greatly extended shelf-life. Due to differences in ionic composition and pH, gel patterns obtained with Bis-Tris gels cannot be compared to those obtained with Tris-glycine gels. Tris-acetate gels are designed for separating large molecular weight proteins and may be used with both SDS-PAGE and native PAGE running buffers. Compared with Tris-glycine gels, Tris-acetate gels have a lower pH, which enhances the stability of these gels and minimizes protein modifications, resulting in sharper bands. The Tris-Tricine gel is a modification of the Tris-glycine gel and is optimized to resolve low molecular weight proteins (2–20 kDa).
The proteins have to work through the mesh structure of the gel as they run. Larger proteins take longer to navigate through the gel, while smaller proteins move faster. The pore size of the gel can be adjusted by changing the percentage of acrylamide in the gel. Less acrylamide means a gel with larger pores, which is optimal for large proteins. More acrylamide means a gel with smaller pores, which is used to resolve smaller proteins. "Gradient gels" are specially prepared to have a low percentage of acrylamide at the top (beginning of sample path) and a high percentage at the bottom (end), enabling a broader range of protein sizes to be separated. Table 2 demonstrates recommended gel percentage based on molecular weight of protein.
|Protein Size (KD)||Acrylamide Percentage (%)|
Table 2: Recommended acrylamide percentage based on molecular weight of protein
Protein samples are separated on gel matrix using electricity by a process called polyacrylamide gel electrophoresis (PAGE). There are several forms of PAGE, with the SDS-PAGE being the most widely used electrophoresis technique separating proteins, which have been resolved into polypeptides, primarily by molecular weight. On application of current, all negatively charged SDS bound proteins migrate through the gel toward the positively charged electrode. Proteins with less mass travel more quickly through the gel than those with greater mass because of the sieving effect of the gel matrix. In native PAGE, proteins are separated according to the net charge, size, and shape of their native structure. Because no denaturants are used in native PAGE, subunit interactions within a multimeric protein are generally retained and information can be gained about the quaternary structure. In addition, some proteins retain their enzymatic activity (function) following separation by native PAGE. Thus, this technique may be used for preparation of purified, active proteins. In two-dimensional (2D) PAGE, proteins are separated by native isoelectric point in the first dimension and by mass in the second dimension.
Traditionally, scientists poured their own gels (handcast) following standard recipes using acrylamide and bis-acrylamide solution, and pouring the mix between two glass plates to polymerize. Gel polymerization in handcast gels occurs in 2 stages. A “resolving” gel is poured first and allowed to polymerize. A “stacking gel,” which has a lower concentration of acrylamide (e.g., 7% for larger pore size), lower pH (e.g., 6.8), and a different ionic content, is then cast over the top of the resolving gel, and the samples are loaded onto the stacking gel once it has polymerized. This allows the proteins in a loaded sample to be concentrated into one tight band during the first few minutes of electrophoresis before entering the resolving portion of a gel.
However, acrylamide and bis-acrylamide are neurotoxins when in solution, so utmost care needs to be observed to avoid direct contact with the solutions and to clean up any spills. Although handcasting offers the benefit of customized gel formulations, the casting process requires hours to complete, and once the gel is polymerized it gets degraded within a couple of days. Handcast gels are thus inconsistent and low in quality. Precast gels on the other hand, offer great convenience, have stringent quality control, higher reproducibility, and longer shelf life than handcast gels. Although expensive, they can be customized per requirement and come in diverse sizes, thicknesses, and formulations to suit customer needs.
Whether using handcast or precast gel, the cassette is vertically mounted into a gel apparatus so that the top and bottom edges are in contact with buffer chambers containing a cathode and an anode, respectively. Gel boxes should be selected based on the size of the gel; commercial gels are usually available as mini or midi gels and have complementary gel boxes. The running buffer is added, which contains ions that conduct current through the gel.
Tris-Glycine SDS Running Buffer (10X) #4050 from Cell Signaling Technology is a 10X stock solution, which must be diluted to 1X before use.
Equal amounts of protein samples are added into individual wells of the gel. A molecular weight marker, which is a premix of several proteins of known molecular masses, is added into a separate well as a frame of reference, and the gel is run for 1-2 hr at 100V. However, some modern gel running equipment can handle running gels at 250-300V for as little as 20-25 minutes. Prestained molecular weight markers #13953 and #14208 or biotinylated ladder #7727 from Cell Signaling Technology cover a broad range of molecular weights. Once the proteins are separated, they can be visualized directly in the gel using Coomassie dye or copper stain.
“Smiling” or “frowning” bands are not uncommon in western blot. This usually occurs when the proteins in the gel are either migrating very fast or if the gel system gets heated up or a combination of both these factors. Optimizing the current and/or reducing the temperature, either by running the gel in a cold room or using an ice-cold apparatus, can help alleviate the problem. Using fresh APS, if using handcast gels, is another way to avoid the “smiling” or “frowning” issue.
Once the proteins have resolved on the gel, they are transferred from the gel onto a sturdy support such as a membrane (nitrocellulose or PVDF) using electric current. Transfer can be done using a wet or semi-dry system. Wet transfer is less prone to failure due to drying of the membrane, and is especially recommended for large proteins. For both kinds of transfer, the membrane is placed next to the gel and should not be touched with bare hands.
Semi-dry transfer: paper > gel
> membrane > paper (all wetted in transfer buffer)
Wet transfer: sponge > paper > gel > membrane > paper > sponge
In semi-dry transfer, the sandwich is placed directly between the positive and negative electrodes. In wet transfer, the sandwich is then placed between positive and negative electrodes and submerged in transfer buffer to which an electrical field is applied. Transfer usually runs at 90V for 1-1.5 hr.
The Tris-Glycine Transfer Buffer (10X) #12539 from Cell Signaling Technology is a 10X stock solution, which can be diluted to 1X before use. Methanol is required to make the 1X buffer.
Two types of membrane are available: nitrocellulose and PVDF (positively charged nylon) depending on experimental conditions.
|Pre-treatment||Needs methanol for equilibration before use.||No equilibration required|
|Protein binding capacity||Has a protein binding capacity of 170 to 200 μg/cm2 and thus recommended for low expressed proteins||Has a protein binding capacity of 80 to 100 μg/cm2 and thus has less sensitivity|
|Binding interaction||Protein binding occurs through hydrophobic and dipole interactions||Protein binding occurs through hydrophobic interactions|
|Binding interaction||Durable and ideal for reprobing||Brittle and not suitable for reprobing|
|Pore size||Typical pore size 0.1, 0.2, 0.45 μm. Smaller proteins require a smaller pore size||Typical pore size 0.1, 0.2, 0.45 μm. Smaller proteins require a smaller pore size|
Table 3: Differences between the two membranes
Air bubbles can hinder the transfer process. It is critical to remove air bubbles in the gel and membrane sandwich by rolling them out using a roller or a pipette, or by assembling the sandwich in transfer buffer to prevent the formation of air bubbles from the get go.
Larger proteins take time to get transferred from gel to membrane, so its best to run the transfer longer. Usually overnight transfer at low voltage of around 35V is ideal for larger proteins (>200 kDa). Adding a small percentage of SDS will help ease the transfer of proteins that may have precipitated in the gel. However, if SDS is added to the transfer buffer, the percentage of methanol in the buffer needs to be lowered as well. For smaller proteins on the other hand, no SDS in the transfer buffer and 20% methanol is more favorable.
Staining the membrane with Ponceau Red aids in the visualization of transferred proteins and is a crude method of determining the success of a transfer. To do this, the membrane should be washed in the washing buffer (1X TBST) after the transfer is complete, followed by staining the membrane with Ponceau Red (diluted 1:100 before use) for 5 min. Once the protein bands start becoming visible, the membrane can be extensively washed in water until the protein bands are clear. The membrane should be destained completely by washing in TBST before continuing the blocking step.
Then Tris Buffered Saline with Tween® 20 (TBST-10X) #9997 from Cell Signaling Technology is a 10X stock solution, which can be diluted to 1X before use.
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 dry milk (#9999) or bovine serum albumin (BSA) (#9998). Milk is cheaper and more popular, but when studying phospho-proteins, BSA is the more popular of the two. To prepare a 5% milk or BSA solution, weigh 5 g per 100 mL 1X TBST. Failure to filter the solution can lead to spotting, where tiny dark grains will contaminate the blot during development. The membrane should be incubated for 1 hr at room temperature with gentle rocking, and washed with 1X TBST 3 times, for 5 min each after the incubation is over.
Primary antibody dilution should be optimized depending on the experiment. Antibodies are usually diluted in blocking buffer at the dilution suggested on antibody datasheet. A high antibody concentration will result in high background due to non-specificity. The time of incubation of primary antibody can vary between a few hours at room temperature to overnight at 4°C, and should be done on a rotator or a rocker so as to keep it agitated. This ensures that the membrane is evenly exposed to the primary antibody. Longer incubations should be done at 4°C to avoid contamination and protein degradation, and to increase antibody binding (Figure 4). After the incubation step, the membrane is washed 3 times with 1X TBST, for 5 min each, to remove residual unbound primary antibody.
A primary antibody should be specific in recognizing the antigen of interest. Both monoclonal and polyclonal antibodies work well for western blotting. Monoclonal antibodies are more specific and reproducible but are more expensive. Polyclonal antibodies are cheaper but lack high specificity and reproducibility. Secondary antibodies must be chosen based on the host species of the primary antibody. If the primary antibody was raised in mouse, the secondary antibody must be anti-mouse. Secondary antibodies can be obtained from several host species, thus offering flexibility to choose the one that gives the least background for your assay.
In general, the primary antibody that recognizes the target protein in a western blot is not directly detectable. Therefore, conjugated secondary antibodies are used as means for ultimately detecting the target protein. Often the secondary antibody is complexed with an enzyme, which when combined with an appropriate substrate, will produce a detectable signal. Considerations when choosing a secondary antibody:
Secondary antibody is usually diluted in 5% Milk in 1X TBST at the suggested dilution. If a titration is required, a range of 1:1000- 1:10,000 should be used to optimize. Secondary antibody incubation is usually recommended for 1-2 hr at room temperature with agitation. After incubating with the secondary antibody, the membrane is then washed with 1X TBST 3 times, for 5 min each, to remove residual unbound secondary antibody.
Detection methods vary according to the customer needs, and often become the rate limiting factors for deciding which route to go. Two standard detection methods in western blot are direct and indirect detection. Direct detection involves using an enzyme- or fluorophore-conjugated primary antibody to detect the antigen of interest. Indirect detection, the more popularly used method, involves using an unconjugated primary antibody to detect the antigen of interest, followed by an enzyme- or flourophore-conjugated secondary antibody to bind to the primary antibody and target antigen complex. Enzyme conjugates include horseradish peroxidase (HRP) or alkaline phosphatase (AP), which when combined with an appropriate substrate, will produce a detectable signal. For fluorescent western blotting, near infrared anti-species IgG conjugates are ideal, e.g., Anti-mouse IgG (H+L) (DyLight™ 680 Conjugate) #5470.
Unlike chemiluminescent or fluorescent blotting applications, detection with chromogenic substrates produces a precipitate on the membrane, resulting in colorimetric changes that are visible to the eye and thus do not require special equipment. However, the colored precipitate formed by chromogenic substrates cannot be easily stripped off to facilitate re-probing procedures.
The most sensitive detection methods use a chemiluminescent substrate that produces light as a byproduct of the reaction with the enzyme conjugated to the antibody (Figure 5). The light output can be captured using film. However, digital imaging instruments based on charge-coupled device (CCD) cameras are becoming popular alternatives to film for capturing chemiluminescent signal.
Alternatively, fluorescently tagged antibodies can be used, which require detection using an instrument capable of capturing the fluorescent signal. Fluorescent blotting is a newer technique and is growing in popularity as it affords the potential to multiplex (detect multiple proteins on a single blot).
Membranes can be stripped of antibodies and reused for the detection of a second protein. A reprobing protocol from Cell Signaling Technology is available online. It is not recommended to strip and re-probe a blot multiple times.
NOTE: The best possible results are always obtained using fresh western blots.Need more help? Watch this easy-to-follow protocol video!