Western blotting, the immunodetection of electrophoretically separated proteins on a membrane, remains the most widely used method of detection and characterization of proteins.

An estimated 80% to 85% of protein papers present Western blot data as part of their findings.

Although the basic principles remain unchanged, Western blot reagents, equipment and best practices have all evolved, sometimes significantly, since the technique’s first description in 1979 [1]. This guide presents information to help beginners with a basic know-how of Western blotting and to inform seasoned users about recent trends and innovations that may improve their workflow, including pros and cons of options currently available in the marketplace.

The Western blotting workflow can be broadly divided into four segments:
1) Electrophoresis or protein separation by polyacrylamide gel electrophoresis (PAGE)
2) Protein transfer from gel to membrane
3) Probing (immunoblotting) of the proteins with labeled antibodies
4) Signal detection

Electrophoresis 

In PAGE, a protein solution travels down a thin slab of polyacrylamide gel under the force of an electric field. Usually, the proteins are solubilized in an anionic detergent (sodium dodecyl sulfate, thus the term SDS-PAGE) that denatures them and imparts a charge proportional to their molecular weight (MW). In a homogeneous gel with appropriate acrylamide concentration, the proteins migrate at a rate proportional to the log of their MW. 

Single-percentage vs. gradient gels

The polyacrylamide (and cross-linker) concentration in the gel determines the pore size, with higher-percentage gels having smaller pores through which the proteins migrate. Lower-density gels are used for resolution of proteins with high MW, and higher-density gels are better for resolution of smaller proteins. Selection charts are available with ranges of recommended concentrations.

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Sometimes a sample contains proteins with a wide range of MWs, in which case a gel with a gradient of polyacrylamide concentrations may be necessary—for instance, when looking for multimers, aggregates and impurities after a single purification step. “This is important because we mostly express proteins for crystallization studies, and for this we need a certain quality of the recombinant proteins,” explains Izolda Popova, assistant director of the Recombinant Protein Production Core at the Chemistry of Life Processes Institute of Northwestern University. “Hence, we need to see everything.”

Selecting the right gel chemistry helps resolve high- or low-molecular weight proteins and may help prevent degradation resulting from, for example, a too low or too high pH. Many such gels, such as tris-glycine, bis-tris and tris-acetate, are available from multiple sources, and others may be unique to a supplier. It is best to check with the vendor to see what gel type it recommends for a specific application.

Precast gels

As a broad generalization, academic labs usually cast their own gels, mainly for cost reasons, and labs in pharmaceutical and biotech companies prefer the convenience and purported reproducibility of (purchased) precast gels. Many tout the superior performance of precast gels. But Popova, who mainly purchases precast mini gels, still prefers the sharpness of the bands in gels she has poured herself, especially for analysis or quality control in very important projects.

Even labs that cast their own homogeneous (single-percentage) gels are likely to purchase precast gradient gels.

The latter are notoriously difficult to pour, even with proper gradient mixing equipment, making gel-to-gel reproducibility more difficult to achieve. Precast large-format gels of either type are generally not available (but casting equipment is readily available), leaving no choice but to pour your own.

Gel boxes

Apparatus for electrophoresis with mini and midi gels (tanks, power supplies and accessories) are available that can accommodate one or more gels simultaneously. Multi-gel boxes typically are configured back to back, requiring the user to reorient the tank (and potentially slosh around buffer) to load the wells. To address this shortcoming, Thermo Fisher Scientific offers a side-by-side gel tank that accommodates two mini gels.

Standard midi gels can accommodate up to 30 µl of sample per lane. For dilute samples, this may necessitate the need to concentrate before loading. Thermo Fisher Scientific offers gels with wedge-shaped wells that allow for up to twice as much sample to be loaded on the same size of gel.

Visit Biocompare product directory to find more on gel boxes. 

Markers

It is standard protocol to dedicate a lane of gel to MW markers. By using pre-stained markers, the lane can serve multiple functions, such as marking the position of a given sized protein on the final blot, visual monitoring of the electrophoretic progress in real time, visual confirmation of protein transfer, definition of blot orientation and matching the type of markers with the way the blot itself is developed—for example, using markers tagged with horseradish peroxidase (HRP) for chemiluminescence blots.

In Bio-Rad’s V3 (“visualize, verify and validate”) system, which uses a Stain-Free™ gel, the gel is activated for 30 seconds following electrophoresis, in a stain-free-enabled imager that links the proteins to small tri-halo compounds resident in the gel. Proteins on the gel (and later, on the membrane) can then be visualized by fluorescence.

Protein transfer 

After the proteins have been run out on a PAGE gel, they need to be transferred to a solid support on which immunoblotting will take place. The most common way to do this is again by electrophoresis,  but instead of the electrical field running the length of the gel, it is perpendicular to the (flat) face of the gel and the adjoining membrane, which are face to face. Proteins migrate onto the membrane in the exact orientation as that on the gel. 

Apparatus

The vast majority of protein transfers take place using wet (also known as submerged or tank) and semi-dry systems. The most typical setup is the wet transfer in which a “transfer sandwich”—consisting of filter paper/gel/membrane/filter paper, surrounded by inert sponge padding and a rigid support—separates two halves of an electroblotting tank filled with a conductive buffer. Wire electrodes in each half of the tank are connected to a power supply, allowing current to pass from one side of the tank to the other through the sandwich, drawing the proteins with it. Wet transfers generally require one to several hours. Because of the amount of heat generated, cooling is necessary; this can be accomplished by a variety of means, such as packing the apparatus in ice, using a heat exchanger or performing the transfer in a cold room at 4°C. Wet transfers require large amounts of buffer, which generally contains 20% methanol; therefore, toxicity and waste disposal must be taken into account.

A semi-dry transfer apparatus is faster than wet and requires far less buffer.

Here, a transfer sandwich of wetted paper/gel/membrane/filter paper is placed directly between two solid plate electrodes. Transfer takes 30 minutes to one hour using standard buffers, but proprietary methanol-free reagents that allow transfer in seven to 10 minutes are also available.

Dry transfer systems work in essentially the same manner as semi-dry systems, although the transfer sandwich (including membrane) are provided as consumables; the gel is just slipped between the layers. The transfer itself takes between three and seven minutes. The Thermo Fisher Scientific iBlot™ 2 and Bio-Rad’s Trans-Blot® Turbo™ are “dry” transfer systems, in the sense that no additional buffer needs to be added.

Wet transfer has generally been considered the most efficient transfer method, especially for higher molecular weight proteins. Manufacturers of some of the newer semi-dry (and dry) transfer equipment claim that this is no longer the case.

Membranes

Membranes for immunoblotting are generally composed either of nitrocellulose or polyvinylidene fluoride (PVDF). Nylon, though popular as a support for nucleic acids in Northern and Southern blotting, is not generally used in Western blotting.

Nitrocellulose membranes are generally slightly less expensive than those made of PVDF. Nitrocellulose is compatible with all detection chemistries typically used for Western blotting and exhibits very little autofluorescence. Yet it is brittle and difficult to handle when dry, making it less than ideal for stripping and re-probing.

PVDF is more robust and has a higher protein-binding capacity, which translates into increased sensitivity. For these reasons, there is a trend toward PVDF as the Western blotting membrane of choice. Yet with higher binding capacity comes higher background. In addition, PVDF is prone to autofluoresce. Low-fluorescence PVDF membranes are available at a premium and are recommended over standard PVDF when using fluorescence detection.

Staining

It’s always a good idea to verify that the protein transfer has taken place. The simplest way to do this is by visual observation of the disappearance of pre-stained MW markers from the gel and their concomitant appearance on the membrane. To verify that all proteins have been successfully transferred, the gel, the membrane or both can be stained. A reversible membrane stain such as Ponceau S, should be used so as not to interfere with the downstream steps. As an alternative, stain-free gels can be used to visualize the total protein. According to a consensus of journal editors [2], total protein staining is the recommended method for normalization of signal intensities in Western blotting.

Immunoblotting: Wash, block and probe 

Western blotting, like other immunodetection protocols, takes advantage of the exquisite specificity of a primary antibody to recognize a particular protein (or epitope) of interest. Generally, the primary antibody is then recognized by another (secondary) antibody specific to it. For example, a primary antibody generated in a mouse binds to protein X, and a secondary antibody, raised in a goat against mouse antibodies, recognizes the primary antibody. The secondary antibody is generally conjugated to an enzyme or a fluorophore, allowing it to be detected downstream. The location of this signal on the blot, relative to the MW markers, allows the size of protein X to be estimated.

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To assure that antibodies do not bind directly to the membrane, it is crucial prior to the addition of each antibody to block any available binding sites with a blocking agent (without displacing the blotted proteins themselves and, of course, the antibodies should not have affinity for the blocking agent itself). The most common classes of blocking agents are nonspecific proteins or mixtures, such as bovine serum albumin, fish gelatin and goat serum; and nonionic detergents. Which blocking agent will work best, and at what concentrations, depends on several factors, including the size of the protein to be detected, the antibodies used and buffer conditions. Several agents are often used in conjunction, and it is worth taking the time to optimize to achieve the best signal-to-noise ratio.

Automated blotting

After the proteins have been transferred to the membrane, the standard Western blotting protocol involves numerous additions, incubations, washes and rinses of the membrane in a rocking/shaking glass or plastic tray. This often totals three hours or more of hands-on time. Several automated solutions on the market are designed to alleviate this tedium, and in some cases they also offer other benefits.

Precision Biosystems’ BlotCycler™, Cytoskeleton’s GOBlot™ and Next Advance’s Freedom Rocker™ BlotBot® offer walkaway blot processing from blocking through secondary antibody incubation and rinsing, while keeping the blot in constant motion. BlotCycler can process up to 12 blots simultaneously and captures the primary antibody for re-use. GOBlot is a smaller, simpler single-blot unit that also recycles primary antibody. BlotBot can process up to six blots and saves both primary and secondary antibodies.

Rather than rocking the blot, Thermo Fisher Scientific’s Invitrogen™ iBind™ uses what the company calls sequential lateral flow technology; this is akin to the capillary forces in paper chromatography to move fluids through the blot without the use of electricity. This method does not recycle the used antibody, but only about one-fifth as much antibody is needed compared with the other manual protocols.

Although manual steps are required for MilliporeSigma’s SNAP i.d.® 2.0 Protein Detection System, it speeds up the entire Western blot processing to 30 minutes by using vacuum power to pull reagents through the blot.

Find more information on automated blotting apparatuses in Biocompare's product directory.

Signal detection

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With radioactivity falling out of favor, there are three principal methods researchers now use to detect the secondary antibody bound to the protein of interest in Western blotting. Colorimetric and chemiluminescent detection rely on an enzyme conjugated to the secondary antibody to catalyze cleavage of a soluble substrate into a colored insoluble precipitate, or the emission of light, respectively. Fluorescence detection typically relies instead on a fluorophore conjugated to the secondary antibody. (Fluorescent products of enzymatic reactions, such as resorufin, can be used for detection, as well.)

Colorimetric and chemiluminescence

There are a host of secondary antibodies conjugated to a variety of enzymes, with HRP being the most favored for Western blotting, followed by alkaline phosphatase (AP). AP has the advantage of a linear reaction rate, allowing for increasing sensitivity with longer incubation times; yet this often leads to high background (and thus lower signal-to-noise ratio). HRP has faster kinetics and better specificity for its substrate than AP, and a larger range of substrates are available for HRP.

Of the nonradioactive Western blotting methods, chemiluminescence (chemi) offers the greatest sensitivity and is the method used by most labs.

It is often called ECL, for “enhanced chemiluminescence,” and its substrates may differ from each other based on the sensitivity of the reagent, sometimes by several orders of magnitude, as well as the duration of the signal. For lowly expressed proteins, a pricier, maximum-sensitivity reagent may be called for. Care in optimization, including perhaps titration of the secondary antibody, may be required because of potential issues with higher background and the appearance of “ghost bands.” Homebrew reagents, based on coumaric acid, luminol and hydrogen peroxide, for example, represent the other option. “We make this solution at large volume, aliquot [it] and freeze it at -20°C. It works perfectly, and it’s cheap,” says Popova.

Unlike colorimetric methods, chemi blots can be stripped and re-probed with a different primary antibody enabling, for example, detection of multiple proteins of nearly the same molecular weight (such as different phosphoforms).

Most labs still read chemi blots by exposing them to X-ray film. In addition to generating hazardous waste, this offers only very limited dynamic range. Digital chemi imagers simultaneously let both weak and strong signals be read and (semi-)quantified.

Read our recent article on western blot documentation systems. 

Fluorescence

Fluorescently labeled antibodies can be detected (with the appropriate reader) immediately after washing off unbound antibody, without the need for additional assay steps. Yet the principal benefits of fluorescence are the ability to multiplex and better quantitation of Western blots.

By using primary antibodies from different species and matching them with secondary antibodies conjugated to different fluorophores, two or three distinct protein epitopes can be simultaneously visualized. To accomplish a similar task with chemi, it would be necessary to strip and re-probe the blot and overlay the results to determine the relative positions of the different signals.

Unlike enzymatic chemi, the signal from fluorescence is linearly proportional to the quantity of antigen on the membrane and to the amount of time it is imaged. The signal should remain stable, and a blot will look the same whether imaged after five minutes, five hours or five days.

Simple innovations

Some alternative technologies are making inroads into the Western market. The explosion of mass spectrometry, for example, gives researchers access to a level of detail about proteins that is unavailable with Western blotting.

Simple Western™ technology from Protein Simple offers automated separation and immunostaining of proteins. But despite the name, “it is not a Western. It’s a capillary-based immunoassay,” points out Joanna Liliental, director of the Translational Applications Service Center (TASC) at Stanford University. Nanoliter amounts of sample are drawn into the capillary, and following size (or charge) separation, a proprietary step uses UV light to covalently attach the proteins to the inside of the capillary. This is followed by the same steps that would take place on a membrane. “After all of this, [a] CCD camera captures the chemiluminescence … and then you have an elegant software that can essentially graph and analyze all your reactions.”

“Quantitation is probably the most important advantage," says Liliental, whose lab uses a 96-sample Peggy Sue™ instrument from Protein Simple. "High sensitivity of detection makes it possible to quantify signal from even picogram amounts of protein. Automation is a big thing, too, because it essentially removes the human error of pipetting,” making the assay simple, reproducible and consistent. A smaller, “more affordable” version called Wes™ processes up to 25 samples, but with lower sensitivity and resolution compared with Peggy Sue, she says. Another option is Protein Simple’s Milo single-cell Western platform. “For people who are looking at single-cell vs. bulk effect, this would answer a lot of questions,” Liliental says. “It’s really cool tech, ” she adds, “but the current iteration is more of a prototype that needs further optimization than a ready-for-prime-time solution.”

LI-COR’s In-Cell Western™ Assay (aka In-Cell ELISA) is designed to interrogate proteins in cells, with the microplate they’re grown in as the solid support. After the cells are fixed and permeabilized, the steps are essentially the same as a standard, multiplex, fluorescent Western workflow.

Conclusion

Innovations and variations likely will continue to make Western blotting faster, easier, more fool-proof, more sensitive, more quantifiable, more automatable and less toxic. But it’s unlikely that the tried-and-true Western blot in its essence will be supplanted any time soon. As the choice of gels, stains, membranes, antibodies, blocking reagents and apparatus available to perform Western blotting continues to grow, users need to think about if—or when—they should upgrade their equipment and protocols. Many factors should be weighed when making such decisions, primarily time and cost. Each option comes with its own advantages and limitations, and user discretion and expertise are vital in choosing the one that best answers the biological question for which the Western blot is to be performed. 

The author would like to acknowledge Karin Söderquist (GE Healthcare Life Sciences) and Priya Rangaraj (Thermo Fisher Scientific) for conversations about Western blotting. Additional information was acquired from GE Healthcare Life Sciences’ “Western Blotting Principles and Methods” handbook, from experts in the topic and from the materials of numerous vendors.

References

[1] Towbin, H, et al., “Electrophoretic transfer of proteins from polyacrylamide gels to nitrocellulose sheets: Procedure and some applications,” Proc Natl Acad Sci USA, 76:4350-4354, 1979. [PMID: 388439]

[2] Fosang, AJ, Colbran, RJ, “Transparency Is the Key to Quality,” J Biol Chem, 290:29692-4, 2015. [PMID: 26657753]