Western Blot Troubleshooting

Common as they are in today’s protein laboratories, Western blots are—let’s face it—a royal pain in the neck.

For one thing, the method itself is lengthy and laborious: run the gel, transfer the gel, block, primary antibody, wash, secondary antibody, wash, wash, wash, detect. Every one of those steps can be optimized, and every one can go wrong.

In fact, some 41% of researchers say their Westerns fail at least a quarter of the time, says Ryan Short, Western blotting product manager at Bio-Rad Laboratories, citing internal marketing data. In other words, it’s not you; Westerns really are that finicky.

Unfortunately, the nature of the protocol also means users rarely realize a given blot is bad until they’ve wasted a lot of time and reagents. “Because Western blotting is something of a black-box process, you don’t know until the very end,” says Short.

One particularly acute problem for Western blotters is sensitivity; it’s easier to detect abundant proteins than those expressed at low levels. But there are things you can try. From blocking buffer to membranes to chemiluminescence substrates, we’ll help you find the formula to make your Western a success.

Western 101

Western blotting is used to detect and quantify (usually relatively) specific proteins in a set of samples. First the proteins are resolved by size on an SDS-polyacrylamide gel and transferred (or blotted) to a porous membrane. The blot is blocked to prevent nonspecific binding and then incubated with an antibody to the protein of interest. The antibody is then removed and the blot washed. Next, a secondary antibody, labeled usually with either an enzyme (e.g., horseradish peroxidase) or a fluorophore, is added, incubated and again washed away.

The final step is detection. With fluorescently tagged secondary antibodies, detection is easy: Simply place the blot in a fluorescence-imaging system, excite at the appropriate wavelength(s) and capture the data. If you used an enzyme-linked secondary instead, you can choose either chromogenic or chemiluminescent detection. The former uses a reagent that forms a colored precipitate on the blot to reveal the bands; the latter generates light, which is detected either on film or digitally.

Chromogenic detection generally is regarded as the least sensitive of the three and chemiluminescence as the most sensitive. “Theoretically you should get as high a sensitivity with fluorescence as chemiluminescence, but we find that ‘chemi’ gives you a better signal,” says David Chimento, assistant laboratory director at Rockland Immunochemicals. “For the most sensitive applications, chemiluminescence is going to get you that data.”

Others, though, say fluorescence vs. chemiluminescence, at least when it comes to sensitivity, is a wash. “I can easily run two assays in parallel, using the appropriate reagents and dyes, which will make fluorescent blotting quite a bit more sensitive than chemiluminescence,” says Dmitry Bochkariov, principal scientist at Advansta, a proteomics tools provider in Menlo Park, Calif.

In other areas, fluorescent Westerns offer a clear advantage. They are more quantitative, for one thing, because the light they produce is directly proportional to the number of fluorophores on the blot, rather than the result of an enzymatic reaction whose output fluctuates, for instance, with enzyme concentration (and thus, protein abundance). Fluorescent blots also are highly stable, meaning they can be imaged at the user’s leisure, whereas chemiluminescence signals decay rapidly.

Most significantly, fluorescent Westerns enable multiplexing. For instance, if a user wanted to detect a particular protein and compare its abundance to a reference housekeeping gene, he could detect those signals simultaneously by coupling two different secondary antibodies to two different fluors. To do the same thing using chemiluminescence, the user would have to detect one signal, strip the blot of antibodies and then repeat the procedure—a method that is tedious and very often incomplete.

Block the noise

One easy way to increase sensitivity is by reducing background. And the easiest way to do that is to optimize your blocking reagent.

“Blocking reagents can have a pretty profound effect on the sensitivity of a blot,” says Short. “Often, this is something that requires optimization and can be specific to the type of sample you’re working on.”

A surprising diversity of blockers is available, and Chimento recommends trying three or four. “It really depends on the primary antibody.” (For instance, avoid nonfat milk-based blockers when probing for phosphoproteins, as milk is loaded with such species, says Deborah Grainger, a product development executive at Proteintech Europe.)

Lindsey Kirby, an applications specialist at Syngene, recommends BSA or SEA BLOCK, a steelhead-salmon extract from Thermo Scientific. LI-COR also offers a nonmammalian blocking solution called Odyssey Buffer, designed primarily for fluorescent applications, as well as a casein-based solution. Users who can’t decide may opt for a Blocking Buffer Sample Pack, which includes sample sizes of both.

EMD Millipore offers its Bløk “noise cancelling reagents” in chemiluminescent, fluorescent and phosphoprotein formulations. According to Brandon McMahon, a research scientist at Millipore, these reagents can be stored at room temperature and use chemicals rather than proteins to reduce unwanted binding.

On a related note, users can also tweak their blocking conditions by changing the concentration of the detergent, Tween-20, from blocking and washing buffers. McMahon suggests removing Tween altogether. But Grainger suggests increasing its concentration from the standard 0.05% to 0.1%. “Tween stops antibodies from getting too ‘sticky,’” Grainger explains, “reducing background signal from any antibody-membrane interactions.”

Enhance the signal

Another variable you can tweak to boost sensitivity, at least with chemiluminescent detection, is the chemiluminescent substrate you use. These reagents often are available in several formulations, which differ in both sensitivity and price.

Millipore’s Luminata Western HRP substrates come in three flavors, Classico, Crescendo and Forte. Classico ($76 for 100 ml) can detect about 6 picograms of GAPDH in a protein lysate, compared with 400 femtograms with Luminata Forte ($109).

Advansta offers its WesternBright HRP substrates in ECL, Quantum and an ultrasensitive Sirius formulation, and Thermo Fisher Scientific’s chemiluminescence substrates come in five formulations: Pierce ECL and ECL Plus and SuperSignal West Pico, Dura and Femto.

According to Priya Rangaraj, market segment manager for protein detection products at Thermo Fisher Scientific, chemiluminescence reagents are not a one-size-fits-all purchase. In other words, you may very well need more than one. Pierce ECL ($131/250 ml) is intended for “everyday use,” whereas the SuperSignal West Femto reagent ($337/100 ml) “can go down to true femtogram-level detection.”

But such reagents aren’t only more expensive. They often produce higher background, too. “With maximum sensitivity comes other problems,” Rangaraj says.

Bio-Rad has attempted to simplify substrate choice with its Clarity™ Western ECL Substrate. As Short puts it, why must users choose between sensitivity and budget? “Choose the affordable [reagent], Clarity, and get high sensitivity at the same time. It’s elegant in that way.”

For those interested in chromogenic Westerns, Thermo Scientific’s Ultra TMB blotting substrate boosts sensitivity to be on par with chemiluminescence, Rangaraj says. And users can get an additional boost with the company’s SuperSignal Western Blot Enhancer. According to Rangaraj, this “hugely popular” reagent (for chemiluminescent, chromogenic and fluorescent Westerns) improves specificity by essentially blocking nonspecific antibody-antigen interactions.

Optimize the blot

One obvious way to strengthen a weak signal is to increase the amount of protein on the gel. But that’s not always possible, Short says, because researchers need to keep the housekeeping protein signal they use for normalization in the assay’s linear range.

Short suggests circumventing that problem by using precast, prestained fluorescent gels (such as Bio-Rad’s Mini-PROTEAN® TGX Stain-Free™ gels, part of the company’s V3 Western Workflow, which includes gels, a blotting system and imager). Basically, Short explains, rather than using a specific protein for normalization, users can use total protein content in the lane instead (which they can record, because the gel is prestained, and the transferred protein is visible on the blot).

“Today researchers basically draw a box around the housekeeping protein and the protein of interest, and normalize,” he says. “We’re saying draw a box around the entire lane and use that data to normalize your protein of interest.”

Because that strategy skips the housekeeping control altogether, Short continues, researchers can max out the protein content in each lane, thereby boosting the signal they care about.

Precast, prestained gels offer other advantages, too, says Short. They are easier and faster than pour-your-own gels, of course, and more reproducible. And they can be imaged before and after use to ensure good transfer to the membrane. (The membrane also can be imaged.) The alternative, Short notes, is Ponceau S staining of the membrane, an option that adds extra steps to the Western process and yet says nothing about how much protein might have been left in the gel.

Speaking of membranes, users have essentially two options, nitrocellulose and PVDF. For most applications, the decision is personal as both work well, says McMahon. However, for fluorescence applications, you’ll want to stick with a low-fluorescence PVDF membrane, as nitrocellulose has high intrinsic fluorescence, says Chimento. If you’re studying small proteins, Grainger recommends Immobilon PSQ PVDF membranes for proteins smaller than 35 kDa. For proteins smaller than 20 kDa, she suggests using a trycine gel to separate your sample.

Finally, make sure you load a good positive control. “If that’s not coming up, then you know there’s something wrong with your antibody,” Grainger says.

Antibody considerations

One final variable to tweak in Western blotting is the reagent at the heart of the process, the primary antibody.

“The Western blotting process is antibody dependent,” Rangaraj says. “So if you have good quality antibodies, you’re going to get good Western blots.”

Look for antibodies that have been validated for use in Western blotting, either by the manufacturer or in the literature, Rangaraj says.

For fluorescence applications, Jon Anderson, senior scientist at LI-COR, suggests the company’s “highly cross-adsorbed” secondary antibodies—that is, antibodies that have been purified to minimize cross-species reactivity, an especially important consideration for two-color fluorescence Westerns.

After you’ve identified the correct antibodies, it’s important to use them properly—that is, at the right concentration. “Users often use too high a concentration, and then you see a lot of background,” Chimento says.

Chimento recommends testing several concentrations of both primary and secondary antibodies to identify optimal conditions. For instance, he says, his team uses some secondary antibodies at a 1:40,000 dilution, far below the “recommended” concentration. At that concentration, he says, “There’s so little molecule that you’re really only binding to the primary [antibody].”

That’s just a taste of the variables users can tweak in Western blotting. If you want more, there are several useful references online, including GE Healthcare’s Western Blotting: Principles and Methods, Thermo Scientific’s Western Blotting Handbook and Troubleshooting Tools and LI-COR’s Good Westerns Gone Bad (available for both chemiluminescence and fluorescence). But before you start running yourself ragged, Grainger offers this advice: “Try the standard protocol first.” You might be surprised.