Expert Coffee Chat Webinar: Western Blotting and Protein Electrophoresis

Adding low concentrations of non-denaturing detergents help solubilize protein aggregates without denaturing the proteins. For best results, use non-ionic or zwitterionic detergents (e.g., Tween 20, CHAPS).

See page 20 of our Electrophoresis Guide for more details:

• A Guide to Polyacrylamide Gel Electrophoresis and Detection

It is likely that heterodimeric protiens will become monomers when subjeted to the treatments in a western blot. Whether dissociation will be complete or not is hard to tell. If you have detection antibodies specifically for monomer or for dimer form, try them sequentially to see what the outcome is. You may be able to modify the sample preparation process to minimize the reduction of the protien.

If you perform a serial dilution of one of your sample and run it on the protein assay kit and spec it, is it linear? If so, then the spec would seem to be okay. If the readings are also off, then there is likely an issue with your spectrophotometer.

When you say your GAPDH is always off, how did you determine that it is off? It is off because you see lane to lane variation on your western blot result but you loaded the same amount of protein into each well in your gel? If so, it can be due to a lot of things: loading variation, uneven transfer of proteins, or western blot preparation. You may be a candidate for stain-free western blotting; using total protein normalization, results would not be dependent on GAPDH or any housekeeping protein.

Learn more about Total Protein Normalization:

• Learning Center: Total Protein Normalization

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If you're seeing smiley faces, first verify that the buffer is loaded correctly, and you may want to try a gradient gel. If the smiling is intermittent, it may be due to gel casting variability; precast gels are generally more uniform than handcast gels, plus they save you time and labor. One thing to look at: is the gel as a whole smiling, or are the bands in each lane smiling. If you're seeing the entire gel smiling, it could be an indication that the voltage is too high; running the gel too fast and overheating.

If you're seeing smiling in each of your individual bands or lanes, then that could be indication of sample overloading. Another cause of smiling can be a difference in ionic strength of your sample solution and the running buffer. If the ionic strength of your sample is too strong, this creates a gradient in the space between your sample lane and the spacers, causing proteins to migrate horizontally as well as vertically. Consistent sample loading across lanes is also important; determine the protein concentration in your samples before loading using a Bradford or BCA assay and adjust as needed to ensure even loading from well to well. One overloaded well will run differently than the well right next to it If you have empty lanes, fill the wells with the same volume of 1X loading buffer.

See the Troubleshooting section of our Electrophoresis Guide for more tips.

• A Guide to Polyacrylamide Gel Electrophoresis and Detection

Yes, tricine gels have been tested with our Trans-Blot Turbo and transfer well. It works well with our standard Trans Blot Turbo packs. I would recommend the smaller pore size, 0.22 um, in either nitrocellulose or PVDF.

• Trans-Blot Turbo Mini 0.2 µm PVDF Transfer Packs

The safest method would be to use precast gels. The acrylamide monomer is the toxic component; once it has polymerized, it is safe. If you have concerns around acrylamide powder in your lab, and purchasing pre-cast gels is not an option, then prepare solutions by performing all mass measurements under the hood.

Please see our selection of precase protein gels.

• Bio-Rad Protein Gels

When your gel begins to smile, it is typically indicative of one of these things:

• Gel became overheated during the run.
  • Check buffer composition (make sure pH not adjusted with acid or base)
  • Ensure buffer is mixed well
  • Check voltage or lower voltage used to run the gel
• Running buffer incorrectly made or reused; use fresh running buffer for each gel.
• Insufficient buffer in tank; fill tank to recommended levels.

Heat can also lead to nonlinear band separation. If the bands continue to merge even after you follow recommendations above, you may run a test gel with only protein standard in every other well to eliminate your sample as a variable to bands merging.

Please visit our Western Blotting troubleshooting guide for more help:

• Troubleshooting Western Blots with the Western Blot Doctor

The voltage that you run your gel is dependent on what type of gel it is. It is best to consult the manual from the manufacturer of your gels. For example, Bio-Rad TGX gels are typically run at 200 V for about 30 minutes but can be run at 300 V if you desire a more rapid run time, with very little loss in resolution, see our quick start guide for details.

• Mini-PROTEAN TGX Precast Gels Quick Start Guide

To resolve issues with lane and band shape or morphology, best practice is to match concentration of protein and salt, and to load equal volumes across the gel. For example, lanes that are blank next to a lane with a high protein load can cause the heavily loaded lane to spread out and become more diffuse as it runs down the gel.

For more troubleshooting tips, see Bio-Rad's Western Blot Doctor.

• Troubleshooting Western Blots with the Western Blot Doctor

There are two common reasons wide/narrow bands occur on a gel. The first is when load volumes differ drastically between lanes. You can make up lower volumes with sample buffer and if you have a blank lane, just load sample buffer to an equal volume. The other reason may be due to salt concentrations being very high or low--in this case, sometimes diluting in sample buffer can help. If salt concentrations are very far off, then buffer exchange will have to be done.

See our Western Blotting troubleshooting guide for more help:

• Troubleshooting Western Blots with the Western Blot Doctor

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It's crucial to quantitate total protein, and Ponceau stain is a popular option. Bio-Rad has a faster alternative, stain-free imaging. So yes, measuring the total protein signal is important; in fact, a number of journals are strongly recommending that as the standard basis for normalization rather than using traditional housekeeping proteins. With Ponceau, the best strategy is removing the stain prior to adding the blocker. The wash buffer, in our experience is really strong. So there's there isn't necessarily a protocol that you could follow. The biggest variable is time, so you need to look at your blot and watch the purple color get removed, with the caveat that in the case with stripping and reprobing, it's difficult to quantify the degree that you've removed your target protein from the membrane also.

If you haven't yet investigated our alternative, we would recommend that you investigate stain-free imaging as the normalization basis, because it does not involve a stripping step or washing step. Regarding stripping and reprobing, our observations with stripping solutions and Ponceau wash solutions is that you can remove from 30 percent up to 70 percent of your protein with each wash cycle, so you run the risk of removing significant amount of your target before you've even had the opportunity to immunoprobe it.

Total protein staining serves as a checkpoint; you can follow the rest of the protocol of the Western blot perfectly, but if your proteins didn't transfer to your blot, everything you do after that is wasted effort. A total protein stain is absolutely critical to verify that you've transferred your proteins efficiently. Our stain-free total protein imaging allows you to do that very easily; you can track your proteins through the entire process, it's a great tool to use.

If your motivation to strip and reprobe is to look for more than one target, then if your stripping reduces the amount of protein you won't get a very accurate quantification. So the other strategy you could look into is using multiplex Western blotting, which can be achieved using fluorescent imaging. So we usually try to refrain from either using Ponceau or stripping and reprobing.

See our Total Protein Normalization page to learn more:

• Learning Center: Total Protein Normalization

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A 16–22 kDa protein is relatively small, but still definitly transferable onto your membrane. Shorter transfer time and smaller-pore membrane are good approaches. If your using semidry transfer, be careful to avoid overheating, which can degrade some of your protein and cause smearing; this can happen if the buffer is not right. You can alos try switching from a cellulose membrane to PVDF or vice versa. You can also try running a gradient gel, for example, a 4–16% or a 4–20%; the denser part of the gel at the bottom may give you a better transfer for those smaller molecular weights. Also make sure you're not overloading those samples because that often leads to really smeary blots.

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It's good that you're checking how much protein is left in your gel post-transfer; this is a crucial quality-control step. The first questions to ask is what are your target size proteins? For proteins >200–300 kDa, consider a wet transfer, and run it slow and long. If your proteins are small but they're not transferring, you may need to check your system; maybe the buffer is not set up correctly.

In addition to optimizing buffer and trying gradient gel, if you're using a nitrocellulose membrane, using a PVDF membranes might also help capture small proteins better. You can also try adjusting the transfer buffer; if you're using a traditional transfer, like Towbin, adding a little bit more methanol helps the proteins to come out of your gel.

See our Protein Blotting Guide for more tips:

• Bio-Rad Protein Blotting Guide

The best blocking buffer is the one that we will give you the least amount of background. However there are blocking buffers to avoid depending on your experiments. For example, if you are interested in looking at phosphorylated proteins, then you want to avoid blocking buffers with casein because phospho-antibodies can react with it. As alternatives, fish gelatin, milk, and BSA all generally work well as long as you use fresh stock and block for the right amount of time. If you are seeing high background in your blots despite trying different buffers, we recommend trying a commercial-grade blocking solution such as EveryBlot Blocking Buffer.

• EveryBlot Blocking Buffer

There is no exact procedure for selecting the best buffer for protein transfer. It is important to determine if you will be using a semi-dry or wet transfer technique and based on that, select the appropriate transfer buffer. If you find that your protein is not transferring well or if you are seeing incomplete transfer, then tweak the duration of the transfer. If that has no effect, then you can slightly modify the transfer buffer formulation. The two most important components of a transfer buffer are methanol and SDS.

Methanol is included in most transfer buffer formulations because methanol aids in stripping the SDS from proteins from separation by SDS-PAGE, increasing their ability to bind to support membranes.

Adding SDS (up to 0.1%) to the transfer buffer increases the transfer efficiency of proteins, but reduces the amount of binding to the membrane. Therefore, if SDS is added to the transfer buffer, it is important to also include methanol (10–20%).

See our Protein Blotting Guide for more tips:

• Bio-Rad Protein Blotting Guide

Both types of membranes are suitable for western blotting but there are some differences between the two.-PVDF has a higher binding capacity so is favored if large quantities of protein are to be loaded. However, for western blotting, this does not have a practical advantage as such high protein amounts cannot usually be loaded in a single lane.

For chemiluminescence detection, both membranes offer equal performance, but for fluorescence detection, Low Fluorescence PVDF has the lowest autofluorescent background. Nitrocellulose has a moderate amount of autofluorescence, while regular PVDF shows high autofluorescence and should not be used. In terms of handling, PVDF is more physically robust while nitrocellulose is fragile. Also, PVDF requires wetting in alcohol to activate it before equilibrating in aqueous buffer while nitrocellulose does not require wetting.

For more information, see our Bio-Rad Protein Blotting Guide:

• Bio-Rad Protein Blotting Guide

Ponceau S is very labile on membranes, and the stain should come off with a few minutes of washing in water. Several water changes are often required to avoid the pinkish hue that remains. If you are finding that the stain is being stubborn, you can wash in TBST for a few minutes, as the added Tween will help the destaining process.

Absolutely. This is one of the advantages of stain-free western blotting. Keep in mind that in order to assess the amount of protein on the gel before and after blotting, you will need to match the characteristics of the two images side by side by selecting the [Copy Transform] button in ImageLab software.

Learn more about stain-free imaging technology:

• Stain-Free Imaging Technology

Smaller proteins will transfer faster than large proteins. It is not uncommon to see some higher-MW proteins remain in the gel after transfer. The main point is knowing the size of your target proteins that is of interest. In order to achieve a more uniform transfer, you may try to run a gradient gel. Use a 4–15% or 4–20% gel for best results. When run using the recommended conditions, these gels leave big proteins in more porous parts of the matrix, allowing for better transfer out of the gel.

Also remember that if you over-transfer in an attempt to get the larger proteins onto the membrane, there is a good chance that your smaller proteins will blow through the membrane, resulting in less yield. So transfer is about balance and having an idea of what you are looking for and choosing conditions to meet those needs.

For more information, see our Bio-Rad Protein Blotting Guide:

• Bio-Rad Protein Blotting Guide

To help choose the optimal membrane for your assay, see Chapter 3 of our Protein Blotting Guide: Membranes and Transfer Buffers (page 18). This transfer guide does a very good job of explaining the differences between types of membranes and which to use for your assay.

• Bio-Rad Protein Blotting Guide

If you are working with a large MW protein (250 kDa and above), check your gel after transfer to determine how much protein remains in the gel after transfer onto the membrane.

PVDF membranes typically have high binding affinity compared to nitrocellulose, but it may be worth trying nitrocellulose because its binding capacity is typically higher. This also depends on the pore size, and optimal pore size is determined by the size of the protein of interest.

When using our Trans-Blot Turbo rapid transfer system, see this document for tips to get a good transfer of large proteins:

• Transferring High-Molecular-Weight Proteins on the Trans-Blot Turbo Transfer System

Avoiding bubbles is always critical to getting a good blot. This is especially challenge when doing tank blots since you have to build the sandwich submerged in the buffer. In semi-dry techniques, this is easier as the components do not float away during assembly. regardless of technique, it is best to roll between each part of the sandwich--roll when you put the membrane on the paper, then again when you put the gel on the membrane, and lastly when you put the last layer of paper on the stack.

Depending on the size of the gel or type of semi-dry device, the best thing would be to follow the manufacturer's instructions. In practice, the semi-dry method is somewhat wet, similar to a wet sponge, but not dripping wet. You should also use a gel roller to roll air bubbles out every step.

For more information, see our Bio-Rad Protein Blotting Guide:

• Bio-Rad Protein Blotting Guide

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How many times you can use the wash incubation solution for a primary antibody depends on a lot of things. It's really something that you'll need to try out empirically. Typically, when you store a primary antibody you want to put it in a solution with a little bit of sodium azide so that nothing can grow in that solution over time. I used to reuse a lot of my housekeeping antibody solutions. I'd keep them in the fridge just as a backup just so I wasn't constantly diluting antibody. I think the best practice, If you're really worried about it, is to just dilute fresh antibody every time, but maybe keep a few dilutions around, especially those antibodies that you know work really well. Maybe keep those in the fridge for a little while until you need them, maybe reuse them a few times.

The primary antibody is perhaps the most important thing in the entire Western blotting assay; it could make or break your experiment, So I would say be very careful if you are going to reuse the antibody. I'm guessing that a lot of the drive to reuse the antibody is for economic reasons, to save money; unfortunately, the type of antibodies that are easier to reuse are the types that identify proteins that are present in high abundance, or the antibody is really very good or very specific. Housekeeping protein antibodies, for example, but those are easily available; almost all the suppliers have them in good quality, and that's why they are not the most expensive ones. The ones that you really want to reuse are those against your target protein, which is likely to be present in low abundance. However, as you reuse your antibody, it loses its potency and specificity each time, so you want to refrain from doing that, especially for target proteins that are present in low abundance, or if the antibody that you have is not a great quality to begin with. So that's a trade-off you'll have to consider when you try to reuse antibodies.

Check out our selection of validated and flourescent western blotting antibodies:

• Western Blotting Antibodies

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If an antibody is showing more than one band, it would be necessary to confirm that in addition to your target protein it's recognizing another protein. If you suspect that it could be from the secondary antibody due to nonspecific binding to another protein, you can troubleshoot that by doing a singleplex of that target protein only and maybe try chemiluminescent detection in parallel with your fluorescent detection, if you have the necessary reagents and imaging system. Buffer problems could cause background, but if you're clearly seeing bands that are nonspecific it's probably not coming from the buffer.

A positive control would be helpful for troubleshooting, if can you find another source for your protein, maybe a recombinant protein. You could run cell lysate to see if hte antibody you're using lights up the same bands or a different band. You could try a different commercially available primary antibody, and run it side-by-side to see if it lights up the same or different bands. It's possibile there is some biological cause from your cells that you don't know about; a band lighting up at the wrong molecular weight could be because you've enriched for a certain species of that protein or maybe it's getting clipped or processed in some way, so another antibody might resolve that. Cell lysis procedures can cause proteins to get clipped; use good protease inhibitors.

You can try different primary antibodies and optimize the detection; most antibody suppliers should provide validation data, so see if that matches the bands you're seeing and if not, call the supplier. Most suppliers have a refund policy or replacement policy if it's not working, and if they don't have validation data for Western blotting, you can call and ask for a sample before you make a purchase. You could also try swapping out the secondary fluorescent antibody with a chemiluminescent secondary and see if the target shows up at the same molecular weight. Another test would be just a no-primary control just to make sure that your secondary is not binding to something nonspecifically.

Our PrecisionAb Validated Western Blotting Antibodies provide validation data including blot images for up to 12 different cell lysates with endogenous protein levels.

• PrecisionAb Validated Western Blotting Antibodies

Protein loss is one of the inherent drawbacks to stripping and reprobing. We recommend starting with the protocol in our Protein Blotting Guide. Stripping is a process that removes bound antibody but unfortunately also will always remove some bound protein. You can adjust the protocol by adjusting the time the membrane is in stripping buffer to minimize loss of bound target while still removing bound antibody. Just make sure to confirm removal of the first primary antibody (incubate with secondary only to ensure you get no signal from the first primary).

Also, make sure to detect the faintest signal first (for example, a protein with low expression) and leave the stronger signals like housekeeping proteins for after the blot has been stripped. Alternatively, if it is an option, you can use fluorescent detection to detect both simultaneously and using different colors to distinguish between targets.

See page 54 of the Protein Blotting Guide for details:

• Bio-Rad Protein Blotting Guide

If doublet bands that are both being detected by the antibody on a blot, this can indicate a number of things: The antibody can be detecting two isoforms of the same protein. For example, if the protein is phosphorylated, it will be slightly larger and run slower in the gel while the unmodified form will form a band below it.

The antibody can be detecting two related proteins. For example, two proteins in a family can share enough sequence identity or similarity to both be recognized. To be confident in your results, you'll need to know that your antibody is specific, and what it is specific to. All the antibody is reporting is that it is sticking to something, roughly how much or how tightly, and the approximate molecular weight of that target.

If you have already performed WB with your proteins and obtained good results, then just keep in mind that the Rf value might be different than the nominal MW of your protein. If you have so far not observed good results and if you are not concerned about the glycosylation pattern, you can enzymatically pre-treat the proteins with Endouncluease A and/or PNGase F. The best practice would be to run samples in duplicate on the same blot (+/- enzyme) to see if any antibody binding has been compromised. If you do care about the glycosylation pattern, then you would need to run with anti-glycan specific antibodies, and ideally in conjunction with other analytical techniques, such as mass spectrometry.

This issue highlights the value of using stain-free imaging as a substitute for traditional housekeeping proteins such as actin. With stain-free gels and stain-free-enabled imagers, you can image the total protein signal prior to any immunodetection step. If you are still using film, then I would suggest immunodetecting your target protein (or proteins) first, then strip and reprobe with actin as the very last step.

Learn more about Total Protein Normalization:

• Learning Center: Total Protein Normalization

Yes; in multiplexing western blotting experiments, multiple primary antibodies are frequently used simultaneously; each targeting one protein of interest. The host species are different so that the secondary antibody will target a specific host species to give you multiple target/channel detection. As far as specificity and sensitivity, as long as your antibody is specific for a target protein, it should not interfere with other antibodies. However, as with every multiplexing experiment, an initial titration experiment will need to be done to know how much of each antibody to use to give you the best result.

Fluorescent Western Blotting Antibodies are ideal for multiplexing:

• Fluorescent Western Blotting Antibodies

If your primary antibodies are high quality, then yes, you should be able to do that. You will know after your first attempt whether there is any non-specific cross-reactivity (just compare the two blots). If they are, then the next step to help with all of the manual steps would be to do multiplex fluorescence, and you can incubate your primaries and secondaries at the same time.

You may wish to explore the capabilities of multiplexed fluorescent western blotting.

• Fluorescent Western Blotting

Bio-Rad recommends starting with the protocol outlined in our Protein Blotting Guide. Stripping is a process that removes bound antibody but unfortunately also will always remove some bound protein. You can adjust the protocol by increasing the time the membrane is in stripping buffer or by repeating the stripping step to remove more bound antibody. For detecting multiple targets, make sure to detect the faintest signal first (for example, a protein with low expression) and leave the stronger signals like housekeeping proteins for after the blot has been stripped.

See page 54 of the Protein Blotting Guide for details:

• Bio-Rad Protein Blotting Guide

A good control would be to spike your purified protein into the cell lysate at a concentration that works when using the pure protein alone. If you can detect it in this case, then it is because the endogenous levels of the protein in the lysate are too low. To maximize sensitivity, you can try using a more sensitive chemiluminescent substrate or moving to a longer wavelength near IR channel if using fluorescence.

Another possibility is that the protein may be blowing through the membrane. Small proteins like these migrate very quickly out the gel and through a membrane. You can test this by re-doing a transfer with purified protein, but add a second membrane behind the first to capture any of this blow through. Then by comparing the signal from the first and second membranes, you can get an idea if any protein is blowing through.

See our Protein Blotting Guide for more tips:

• Bio-Rad Protein Blotting Guide

The proteins that you have listed are all 35–55 kDa. Depending on what sample type you are using, some other alternatives are vinculin (125 kDa) for whole cell or transferrin (75 kDa) for serum. Alternatively, you may want to explore stain-free western blotting; using total protein normalization, your results would not be dependent on any housekeeping protein.

Learn more about Total Protein Normalization:

• Learning Center: Total Protein Normalization

The typical concentration of Tween in a wash buffer is 0.1%. If you need to make adjustments because of excessive background, I have seen researchers go up to 0.5% Tween for the wash buffer. Any higher may be on the extreme end. For antibodies, I would stay at 0.1%. If you are still getting high background, your antibodies may be binding unspecifically.

See our Protein Blotting Guide for more tips:

• Bio-Rad Protein Blotting Guide

There can be many factors making this protein hard to detect. Is there an well established antibody for this protein or is this a novel protein? How is the stability of the protein when denatured? It is in very low abundance in your sample? Is there some biology going on which makes it hard to detect?

The most important point is to understand the questions above to know what the limitations are before you begin your experiments. If the protein is unstable when taken out of its physiological state, then you may want to find a way to stabilize it, change buffer conditions, express it as a fusion protein if possible. If it is in low abundance, then consider enriching your sample by immunoprecipitation.

If you want to reuse secondary antibodies, the most important thing is to minimize cross-contamination. It is very easy to wash off some of the first primary and introduce it to the secondary antibody solution. This would lead to unexpected bands when the secondary is reused. Additionally, be aware that once diluted, the HRP that is conjugated to the secondary may become less active, reducing sensitivity. Most people do not reuse secondary antibodies because they tend to be relatively inexpensive (compared to primary anitbodies).

Re-incubating a blot with a more sensitive substrate is a commonly used trick. If you do this, be aware that some formulations of substrate may not be compatible with each other. Best practice is to do a wash with TBS.

Carryover is one of the inherent problems with stripping and reprobing. Since you are detecting two very strong targets, you can strip more thoroughly without risk of losing too much protein. You can strip twice or just leave the membrane in the stripping solution for a longer period of time. Just make sure to confirm removal of the first primary antibody (incubate with secondary only to ensure you get no signal from the first primary).

Alternatively, if it is an option, you can use fluorescent detection to detect both simultaneously and using different colors to distinguish between targets.

Learn more about multiplex fluorescent western blotting:

• Fluorescent Western Blotting

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This debate has been around as long as digital imaging technology has been around - about 20 years. A few years ago it was probably true that film imaging was more sensitive than digital imaging for Western blotting, especially in the beginning of the technology, but digital imaging has since improved. And there are other benefits users get by switching to digital imaging for Western blotting; for example, your data are created in a digital format, so it's easy to analyze data, especially if you're doing quantification, as in the previous question. It's easy to manage and archive data, and obviously you can print it as well and get a hard copy, but with the film you start with a hard copy which needs to be scanned if you need to analyze it. Also, because film is so sensitive, it's difficult to manage the exposure, especially if you have a protein that is present in high abundance, like a housekeeping protein, Gapdh, for example, it will saturate in a fraction of a second, and you don't want that. To have quantified data, you need a linear signal, and you cannot get that if one of your signals, especially your control, is oversaturated.

Regarding sensitivity, that may have been true a few years ago, but digital imaging technology has come a long way, and now we're seeing there are few imaging systems on the market, especially from some of the manufacturers that have been around for a long time, that are actually now as sensitive, or more sensitive, than film imaging for chemiluminescent Western blotting. Another advantage of digital imaging is you have the option of switching to fluorescent Western blotting, where obviously the biggest advantage is the ability do multiplex blots and rad imaging, so you can identify and also quantify more than one protein on the same blot without having to do strip and reprobe. In summary, while film was more sensitive than digital imaging in the past, that's not true now, and digital imaging provides you with so many other benefits including the ability to do both chemiluminescent and multiplex fluorescent western blotting.

See our Film vs. Digital Western Blot Imaging page to learn more.

• Film vs. Digital Western Blot Imaging

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Here again, the problem is to identify and quantify more than one protein from the same gel. Strip and reprobe is one strategy, and cutting the membrane is another strategy. We recommend to avoid both methods and instead use and alternate strategy, for example, multiplex fluorescent western blotting, or using a total protein normalization, either Ponceau stain or the recommended technology, strain-free imaging However, if these options are not available, then cutting the membrane is better than stripping and reprobing. You should definitely image them at the same time so you have the same exposure on both; keep them side-by-side so you expose them for the same amount of time. It's best to use a digital imager that has a wider dynamic range, which will help you to capture both the target protein, which is usually present in much lower quantity on your membrane than the protein used for normalization, such as Gapdh. An imaging system that has a wide dynamic range allows you to remain within the saturation limit of your housekeeping protein.

As for incubating them together, because you're probing them with different primary antibodies, you'll need to incubate them separately with different antibodies, which is not the best way to do it. However, you could incubate together when multiplexing, because you'll be probing both proteins using different secondary antibodies, and those two secondary antibodies would be visualized using different conjugated dyes, which minimize the crosstalk. But if you're using chemiluminescent detection on both, you'll have to incubate them with the primary antibodies separately.

One one thing we want to caution you about is related to journal publication standards; for example, the Journal of Biological Chemistry has a number of requirements for submitting blotting images; one requirement is that with your target and your housekeeping bands, your image shows at least two bands of the ladder, one each of molecular weight below and above your housekeeping band. This may not be an issue if your housekeeping protein is a significantly different molecular weight than your target, but keep in mind that this is a requisite for submission, and so you might encounter issues if you were to cut it and you weren't able to show two standards on each strip that you cut.

Learn more about recent changes to journal submission requirements:

• New Publication Requirements for Quantitative Western Blotting

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Background arises from many sources, and every step of the western blotting workflow can contribute to background. Blocking is an important step, and washing steps are very important, but you may need to determine exactly where your background is coming from, or identify what type of background you're seeing and diagnose. For example, a splotchy background all over the blot could be an indication of incomplete mixing of your buffers, or when you're washing your membrane, the agitation of your buffer is not uniform. If you're seeing a more evenly distributed background, slightly hazy or gray everywhere, then that could be an indication of antibodies not binding specifically. So it may require some troubleshooting to determine the source of high background.

For specific tips on dozens of specific western blotting problems, see our online help guide:

• Troubleshooting Western Blots with the Western Blot Doctor

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This is a big problem using darkroom film development for chemiluminescent imaging. With digital imaging, you have more control over the exposure because it does not saturate so quickly, especially for high-abundance proteins. With film, you can get a signal that's already oversaturated within a second, whereas with digital imaging, it might take few seconds, which means you have more control over exposure, and you can get a signal that is just below saturation.

It's easier to see where you're at in terms of saturation with digital imaging because you can see the intensity of the signal directly; for example, using a modern imaging system with a touch screen, immediately after acquisition you can zoom in and you can check the signal intensity at a pixel level. You want to be under 100%, ideally close to 90–95% for a strong signal.

To get both a strong signal and a weak signal on the same blot is challenging, and digital imaging will definitely help you do that as compared to film, because film has a very low dynamic range. With digital imaging systems, especially those a 16-bit algorithm and a camera chip to support that, like Bio-Rad imaging systems, you can capture signals in different ranges, form very low to high-abundance proteins or that are recognized by the primary antibody very well and then the ones that are not.

Another option with digital imaging is multiplex fluorescence. If you think your two signals are very far apart, you can capture them using different channels. For example, on one channel for the target protein that is very low abundance, you can use a very bright fluorophore like infra 800 or 700 on your secondary antibody and let it go for several seconds up to several minutes of exposure if you need to. Then on the second channel, you can use a secondary antibody which is in the blue or green range, away from the infrared, so there is no crosstalk and that image can be captured very quickly within a few seconds, if it's something like a Gapdh.

Another benefit of digital imaging is you can identify the regios of the blot that are subject to saturation. For example, when using chemiluminescence, you can acquire a number of images at varying exposures, perhaps reducing by 10% each time to determine optimal exposure time to obtain a good-quality image without saturation anywhere to yield the most accurate quantification.

Learn more about film vs. digital imaging for western blotting: Film vs. Digital Western Blot Imaging

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Choosing the right housekeeping protein is a challenge because if you're changing biological conditions, our manipulation may affecting housekeeping protein expression levels. It's a good idea to pick multiple housekeeping proteins to test, but maybe a better idea would be to quantitate your total protein and normalize to the total protein in that lane. You can do that with our stain-free imaging technology, and our image lab software makes it very easy. Instead of calculating the ratio of your target signal over your housekeeper, use the ratio of your target signal over total lane protein, and you can compare that with the data you're getting from your housekeepers to ensure that you're getting a good response. Redundancy can help you verfiy that you've chosen the right housekeeper; you can try a few of them. Actin, Gapdh, and tubulin are the most common ones.

See our page on Total Protein Normalization to learn more.

• Learning Center: Total Protein Normalization

The stain-free compound will bind to tryptophan residues in your protein. However, the modification is very minor, so it does not interfere with downstream applications like mass spectrometry analysis of the target protein. In addition, not all Trp residues will be modified. All it takes is one Trp modification to enable detection of the stain-free signal. Furthermore, as long as the antigen-binding site of the antibody you wish to use is not at the Trp molecule, it will not affect detection.

For additional information on stain-free technology, visit our page below.

• Stain-Free Imaging Technology

For a visible marker that also shows up in fluorescence channels, it will depend on what channels you are using. Most people are likely using Red or Far Red as at least one of their channels. In this case, we recommend the All-Blue standard. You can also see our Standards selection tool here:

• Protein Standards Selection Guide

We recommend using total protein normalization (TPN) as the method to normalize your blot for quantification of your western blot.

See our page on Total Protein Normalization to learn more.

• Learning Center: Total Protein Normalization

Ghost bands on film can be caused by moving or repositioning the membrane on the film during exposure. For example, if you first place the blot on the film but need to reposition it, the film will see the blot twice, once in the first position, then once in the second, final postiion. This can lead to ghosting.

See our Western Blotting troubleshooting guide for more help.

• Troubleshooting Western Blots with the Western Blot Doctor

We recommend loading 10–50 ug of total protein sample per lane when imaging stain-free gels for western blotting. This allows sufficient signal for quantitation without overloading. Another suggestion is to use low-fluorescence PVDF membrane to reduce membrane background; nitrocellulose is OK, but regular PVDF has very high background.

For details, see our general Stain-Free Western Blotting protocol:

• Stain-Free Western Workflow: Blotting Protocol

Yes, you can cut the membrane. Make sure that cutting the blot does not prevent you from using the stain-free total protein signal signal as a loading control. One of the major advantages of stain-free blotting is the ability to use total protein normalization as a loading control instead of using housekeeping proteins to normalize your quantification to all the protein in each lane, eliminating the need to cut the blot in most cases.

See our page on Total Protein Normalization to learn more.

• Learning Center: Total Protein Normalization

The propriety fluorogenic compound in our stain-free gels is activated by a UV light source. We recommend using a Bio-Rad Stain-Free enabled imager to capture this signal for total protein normalization. These imagers are designed to yield the best signal from Stain-Free gels and be consistent across gels.

See our page on Total Protein Normalization to learn more about it and how stain-free blotting can help your research:

• Learning Center: Total Protein Normalization

There are strong recommendations by JBC to eliminate the cutting of blots into strips in order to minimize variables/artifacts which may confound your results. This is not to say that no one is doing this anymore. As long as you understand the implications doing so may have on your data, you can take steps to minimize it. If detecting multiple target proteins on the same blot is a must for you, fluorescent western blotting may be something that is worth looking into.

Learn more about the possibilities of fluorescent western blotting:

• Fluorescent Western Blotting

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Yes, western blotting is quantitative, but there are two different types of quantitation methods: absolute quantitation and relative quantitation. With the former, one is able to obtain an actual protein concentration of their target, whereas with the latter, it's relative abundances. The majority of people do relative quantitation because what they're looking for is a regulatory event, whether it's an upregulation or down-regulation compared to their control sample, so that's considered relative quantitation. By contrast, in absolute quantitation, one can obtain an actual protein concentration, but that does require a standard curve within the gel itself, typically three to five points. The disadvantage here is that one has to sacrifice the number of lanes in to generate that standard curve. Be aware that both techniques are available, and depending on what what specific question you're trying to answer, then you can use one of those two techniques accordingly.

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Western Blots are challengning because of the multiple steps involved and the countless variables that can arise from start to finish. Many researchers try to follow a SOP or protocol that has been developed in their lab over time. This does not always work because samples change over time, small variations in gel/buffer formulation can also impact result. We understand this is an issue and our company; over the years; have developed useful tools to steam line the western blotting process while generating reliable data. In addition, we have a fantastic Technical Support team that is very knowledgable in everything western blotting are ready to help. We want you to have a positive experience in the lab and ultimately generate data that helps you move forward with your reasearch.

Learn mor about our streamlined approach to Western Blotting:

• Stain-Free Western Workflow

Single-cell western blotting is an approach in which data is generated from very little protein input. There are various ways to achieve this; one way is by way of a microscope slide with a thin layer of acrylamide.

• Hughes A et al. (2014) Single-cell western blotting. Nat Methods 11, 749–755. DOI: 10.1038/nmeth.2992
• Murphy R (2014) Single-Cell Western Blot and Stain-Free Total Protein Loading Control (webinar)

Glycoprotiens can be tough because depending on the glycosylation state, you may see a smear on your blot. If you have already performed WB with your proteins and obtained good results, then just keep in mind that the Rf value might be different than the nominal MW of your protein. If you have so far not observed good results and if you are not concerned about the glycosylation pattern, you can enzymatically pre-treat the proteins with Endouncluease A and/or PNGase F.

The best practice would be to run samples in duplicate on the same blot (+/- enzyme) to see if any antibody binding has been compromised. If you do care about the glycosylation pattern, then you would need to run with anti-glycan specific antibodies, and ideally in conjunction with other analytical techniques, such as mass spectrometry.

Regarding technical replicates on western blotting and a related question of whether duplicates with varying concentrations add robustness to your interpretation: The best practice for replicates is to use completely different gels and blots. This is to minimize possibilities of artifacts that could affect replicates on the same blot--for example, if a pipetting error was made during antibody solution preparation.

In terms of using different concentrations, the reason that you might want to consider changing concentration is if you are worried that your signals are not within the linear range and one may be saturated. In this case, reducing the concentration could help. Also make sure you load sufficient amounts of sample so you are not hiding unwanted bands. As long as your signal is within the linear range, and above the lower limit of quantification, then there is not much benefit to using varying concentrations of samples.