Protein Stains Support—Troubleshooting

Any staining procedure that uses over 50% alcohol will dehydrate a gel. Simply rehydrate the gel in some water.

The problem is related to the TCA not being rinsed off as it lowers the pH of the solution and causes aggregation of the stain. We recommend washing in large volumes of water, twice for 5 minutes each and performing the last wash for at least an hour (can even go overnight if needed). Another recommendation is to use excess amount of SimplyBlue™ SafeStain.

This is most likely due to SDS interference. We recommend washing the gel extensively before starting the staining procedure.

Here are possible causes and solutions:

Imperial™ Protein Stain contains additives that help slow formation of dye-dye aggregates, which occur in all Coomassie dye-based protein staining reagents. If left undisturbed, the reagent may form visible aggregates that settle in the bottle; however, gentle mixing completely disperses these aggregates. To ensure that a homogeneous sample of the reagent is used, mix reagent before pouring or dispensing.

This is most likely due to SDS interference. We recommend increasing the number of washes and/or wash volumes before staining. Destaining the gel for 5 minutes with 25% isopropanol/10% acetic acid solution or 12% trichloroacetic acid will also help.

Here are possible causes and solutions:

This is most likely due to a problem with the western transfer. Please confirm that the transfer buffer and transfer conditions are correct.

This is most likely due to insufficient destaining time. We recommend destaining the membrane in 30% acetonitrile/20% ethanol solution for an additional 5 mins.

A reduction in background may be seen if the protocol for NuPAGE™ Bis-Tris/Small Peptides is used instead. It uses an extra fixing step to remove excess SDS, which can act as an anti-colloidal agent and cause higher background. The low pH of the staining solution will fix the gel, but not as fast as the pre-fixing step performed in the NuPAGE™ protocol.

• The background is higher in low percentage acrylamide gels due to penetration and trapping of colloids within the large pores of these gels.
• Background may be removed by incubating the gel in 25% methanol solution until a clear background is obtained. Be aware that dye will also be partially removed from the bands.
• Prolonged incubation in >25% methanol results in complete destaining of protein bands and background.

No, the blue "chunks" you see are called colloids. They enable the stain to work effectively. We recommend shaking the solution well before using to evenly distribute these chunks throughout the bottle.

When Colloidal Blue Stainer A and Stainer B solutions are combined, a precipitate may form; this will dissolve within 30 seconds of gentle shaking.

Probably not, however if you add >10% less than the total required amount of Stainer B solution, your gel may not stain as intensely as it would have with the proper amount. Adding more of the Stainer B solution will not affect results, nor will +/– 50% of the Stainer A solution.

Colloids, or blue chunks, are routine in this type of staining procedure. However, if the amount is abnormally high, it usually indicates that too little or no methanol was added.

Yes, you can either completely destain the gel in water and start staining from scratch or if you feel that the staining intensity is a little low and you would like to darken it, you can directly place the gel back in the staining solution.

Background is generally higher in gels less than 10% in acrylamide concentration due to penetration and trapping of colloids within the large pores of the low-percentage acrylamide gels. Excess background may be removed by incubating in 25% methanol solution until a clear background is obtained. Be aware that dye will also be partially removed from the bands and prolonged incubation in >25% methanol will result in a complete destaining of protein bands and background.

Here are possible causes and solutions:

Here are possible causes and solutions:

Here are possible causes and solutions:

Here are possible causes and solutions:

The frozen SilverXpress™ Staining Kit should be fine to use; frozen kits have been tested in the past and have shown to give an equivalent performance.

Here are possible causes and solutions:

• Contaminants from the sample wells entered the gel. Carefully rinse the sample wells with 5 or more changes of 1X running buffer prior to sample loading.
• Poor water quality. Use ultrapure water of >18 megohm/cm resistance for preparing solutions or rinsing.
• Contaminated equipment used to prepare reagents - Use glass columns and sterile pipettes to prepare reagents. Wash glassware thoroughly.

This is probably keratin contamination from fingertips or airborne sources. We recommend wearing gloves at all times during electrophoresis and staining steps, and rinsing the gel wells with ultrapure water or running buffer before sample loading.

Here are possible causes and solutions:

• Poor water quality. Use ultrapure water of >18 megohm/cm resistance for preparing solutions or rinsing.
• Staining trays not clean or containing solutions left over from prior silver staining. Use staining trays dedicated for silver staining. After silver staining, wash trays with soap and water, and rinse them with ultrapure water.
• Improper washing done between steps. Do not skip or reduce any washing steps.
• Gels are bent or torn. Be careful during handling of the gel. Remove the gels carefully from the cassette after electrophoresis making sure that the gels do not tear.
• Gels are not completely submerged during staining. Be sure to completely immerse gels in staining solution and perform all steps using a rotary shaker for even staining.

The length of the soaking interval can be extended if there is suspicion of protein loss through incomplete fixation. However, overnight fixation diminishes stain performance. If fixation times are significantly extended accidentally, then additional post-development washes are recommended to minimize gel cracking during subsequent drying.

No, it is not possible to reverse the process if the gel is overstained. If left in the Developing solution, the gel will continue to darken and then turn black within 30 minutes. It is therefore very important to carefully monitor the development process and add the Stopper reagent at the appropriate time. Since penetration of solutions into the gel is not instantaneous, it might be necessary to add the Stopper reagent slightly before the desired band intensity has been attained. This is especially the case in heavily loaded gels where developing proceeds very rapidly.

If the solution turns brown momentarily and then turns clear, this is normal (sometimes this happens so quickly, that this process is not observed). However, if the solution remains brown, this is an indication of a contaminant in the cylinders used to mix the stains, usually introduced by using the same cylinder that contained a prior solution.

Here are possible causes and solutions:

• Low protein load. Increase the amount of protein loaded. Be sure to have at least of 1–5 ng protein on the gel.
• Poor water quality. Use ultrapure water of >18 megohm/cm resistance for preparing solutions or rinsing.
• Incorrect volumes of water used for rinses. Use exact volumes of all components and strictly adhere to the protocol.
• Stainer or developer solution not prepared properly. Assuming that the sample load is sufficient, the most likely cause for staining failure is improper preparation of either the silver staining solution or the developing solution. If no bands are observed within 5 minutes of adding the Developing solution, we recommend adding 5 mL of the Developer directly to the staining tray. Make sure that the solutions are prepared correctly using ultrapure water.

Thiosulfate, known as "Farmer's Reducer", may be used to attempt to decrease the background. However, it destains the bands as well, so the concentration should be diluted. Leaving the gel in stop solution longer than recommended will decrease background and band intensity, as well.

In general, background staining in Tricine gels is slightly higher than in Tris-Glycine gels. The relatively high concentration of solutes in Tricine gels as compared to their Tris-Glycine counterparts appears to slow the rate of solution exchange into the gel. This can be counteracted by increasing the soak time in the sensitization step (you may leave it in overnight) as per the modified procedure, and then proceeding.

Use the following visual cues as landmarks of a properly completed step:

• The low-percentage stacking gel appears whitish opaque as compared to the separating gel after the Sensitizing Step indicating that this step was performed correctly.
• Mini-gels curl up into a cylinder and float on the surface during the first water wash after the Sensitizing Step.
• When Stainer A and Stainer B are added together, a brown precipitate is formed, which is visible only momentarily. This brown “flash” is a good indicator that the staining solutions are mixed correctly. If the brown color does not revert to clear, discard the solutions, obtain clean glassware, and remix the solutions.

This is usually due to overloading of the protein sample. We recommend decreasing the protein load per band. For an unknown protein, a serial dilution may be necessary to determine the best amount to load for a particular protein.

Here are possible causes and solutions:

• Stopper not added to the gel at the appropriate time. Be sure to add the stopper slightly before the desired stain intensity is reached.
• Protein is overloaded. Decrease the protein load on the gel.

Here are possible causes and solutions:

• High concentration of DTT (>50 mM) in the sample. Use 30–50 mM DTT for reducing the sample.
• To prevent streaking, reduce and alkylate the sample as follows: Reduce the sample with freshly prepared DTT to a final concentration of 17 mM and heat the sample at 70 degrees C for 10 minutes. Then, alkylate the sample with freshly prepared iodoacetamide to a final concentration of 35 mM and heat the sample at 70 degrees C for 10 minutes. Add SDS sample buffer without DTT to the reduced and alkylated sample and proceed with electrophoresis and silver staining.
• Presence of thioflavin in the sensitizer. We recommend heating the 30% ethanol wash in the microwave oven before adding it to the gel after the sensitizing step, and also washing the gel with water a bit longer. You can try this with the turbo method but you risk losing the effects of the sensitizer.

Here are possible causes and solutions:

• Poor water quality. Use ultrapure water of >18 megohm/cm resistance for preparing solutions or rinsing.
• Staining trays not clean or containing solutions left over from prior silver staining. Use staining trays dedicated for silver staining. After silver staining, wash trays with soap and water, and rinse them with ultrapure water.
• Improper washing done between steps. Do not skip or reduce any washing steps.
• Gels are bent or torn. Be careful during handling of the gel. Remove the gels carefully from the cassette after electrophoresis making sure that the gels do not tear.
• Gels are not completely submerged during staining. Be sure to completely immerse gels in staining solution and perform all steps using a rotary shaker for even staining.

Here are possible causes and solutions:

• Loss of silver ions from the gel. Limit the wash after staining to 30–60 seconds.
• Stainer or developer solution not prepared properly. Make sure that the solutions are prepared correctly using ultrapure water.
• Low protein load. Increase the amount of protein load. Be sure to have at least 1–5 ng protein on the gel.

Here are possible causes and solutions:

• Stopper not added to the gel at the appropriate time. Be sure to add the stopper slightly before the desired stain intensity is reached.
• Protein is overloaded. Decrease the protein load on the gel.

This is probably keratin contamination from fingertips or airborne sources. We recommend wearing gloves at all times during electrophoresis and staining steps, and rinsing the gel wells with ultrapure water or running buffer before sample loading.

Here are possible causes and solutions:

• Low protein load. Increase the amount of protein load. Be sure to have at least 1–5 ng protein on the gel.
• Some proteins may need longer fixing time. Increase the time for fixing the gel to 2 hours or overnight.

This is usually due to overloading of the protein sample. We recommend decreasing the protein load per band. For an unknown protein, a serial dilution may be necessary to determine the best amount to load for a particular protein.

Shadowing around the bands means that the gel background staining of SDS is too high. Destain the gel in 10% methanol/7% acetic acid a little longer, approximately another 30 minutes and then give it a good water wash. In the future, try fixing the gel longer, at least another 30 minutes, to better remove SDS from the gel and reduce initial background staining.

SYPRO™ Ruby Protein Gel Stain is not stable beyond about a year. The dye begins to precipitate out from solution (self-aggregate) over time and will show a lower staining intensity of protein bands and increased ‘debris’ or ‘speckles’ on the surface of the gel. It is not possible to filter the stain to remove dye precipitate, as the dye sticks to most paper and membrane filters and will be removed from the staining solution.

Speckles on the gel can increase as the SYPRO™ Ruby Protein Gel Stain ages, due to self-aggregation of the SYPRO™ Ruby dye over time. Speckles can also form due to dye aggregation around contaminants from the staining container, solutions, or particles from the air or gloves, including keratin proteins from skin and hair. When gels are incubated with SYPRO™ Ruby Protein Gel Stain for several hours or longer, dye can build up on the sides of the staining container and then be dislodged with continuing rocking, especially during the destain step, forming speckles. Non-dye speckles can also show up in the image from auto-fluorescent particles of dust, hair, glove powder, or clothing lint that falls on the gel or surface of the glass imaging plate. The better the imager is at focusing on surface features of the gel, the more speckles that are going to be visible.

To minimize the formation of speckles and other background debris, follow clean laboratory practices, use ultrapure water of >18 megohm-cm resistance to prepare solutions, rinse gloves in water to remove powder residue before touching gels, use lint-free wipes and wear a lab coat or avoid wearing clothing that generates a lot of lint, always rinse the staining container with ethanol and wipe out any residual dye before staining another gel, and always rinse and wipe down the glass imaging surface with ethanol and water before placing your gel down. Remove dye buildup on the surface of the staining dish by wiping out the dish with ethanol between the stain and wash step. The rapid stain protocol is complete in as little as 90 minutes, which does not allow enough time for most speckles for form. Once speckles have been deposited on the gel, it is not possible to wash them off.

Speckles will show up as sharp, tall spikes on 3D renditions of gel images. These spikes look distinct from 3D renditions of protein spots or bands. Some image analysis software packages have de-speckling algorithms that can easily identify and remove this type of pixelated noise.

These types of spots are caused by some component of the IEF sample buffer that runs into the gel during the second dimension separation and is stained by SYPRO™ Ruby Protein Gel Stain. SYPRO™ Ruby dye is attracted to amines, such as amines in lysine, arginine and histidine containing peptides, but also amines in detergents and ampholytes. The simplest solution is to try a different IEF buffer formulation that does not cause this artifact. Possibly, a thorough overnight fixation in several changes of 50% methanol/7% acetic acid will wash out the contaminant.

Blue-colored dyes absorb light in the red wavelengths, so they absorb the red fluorescent emission of SYPRO™ Ruby dye. SYPRO™ Ruby dye still binds these proteins, but the signal is quenched by the colored dye, resulting in a negatively stained, dark band. Examples of molecular weight markers with blue-colored proteins that will quench SYPRO™ Ruby fluorescence are the BenchMark™ Pre-Stained Protein Ladder and some proteins in the SeeBlue™ Plus2 Pre-Stained Standard. The same phenomenon can be seen with the bromophenol blue dye front, if it is not completely run off the gel, and loss of signal when SYPRO™ Ruby stained gels are subsequently stained with Coomassie Blue stains. Most other colored dyes do not quench the SYPRO™ Ruby dye signal and will appear as normally stained protein bands.

Your samples or the gel wells were contaminated with keratins from skin or hair. Rinse out the gel wells with ultrapure water or running buffer before loading samples. Wear a lab coat and gloves when preparing samples and use microfuge tubes that have been stored in sealed plastic bags, not left out on the bench top, for preparing samples.

This is likely due to low amounts or no protein present in the sample. Please determine the protein concentration in the original sample.

Here are possible causes and solutions:

Here are possible causes and solutions:

• High protein concentration on specific membrane site. Reduce protein concentration in the transferred gel.
• In some cases, proteins reduced with DTT may not erase completely. Extend erasing step in the Eraser solution for up to 5 minutes.

This could be due to the presence of detergent. Use clean trays when applying the stain. Use separate trays for downstream processing such as immunodetection.

We recommend completing the erasure step and then re-staining the membrane and proceeding with the protocol.

Here are possible causes and solutions:

• Inadequate staining. Use appropriate staining protocol based on the gel type. Use BenchMark™ His-tagged Protein Standard as a positive control to verify staining reagents and protocol. Avoid excessive washing of the gel.
• The gel is not visualized or imaged properly. Be sure to visualize the gel using a UV transilluminator equipped with a camera or a laser-based scanner using the correct filters (see manual for details). A Polaroid™ camera is not recommended. Make sure the aperture on the camera is open wide to allow enough light entry and that the camera is connected to imaging software that allows contrast adjustment for viewing the best image. Visualize the gel immediately after completing the washing steps. Storing the gel in phosphate buffer decreases the signal intensity.

Low protein load or expression level. Check total protein content of the gel by staining the gel with a total protein stain (check page 13 of the manual). Load at least 1 pmole of the His-tagged fusion protein for detection. Make sure the His-tag is in-frame and the protein is expressed properly.

Here are possible causes and solutions:

• Missed washing steps. Be sure to wash the gel twice with 20 mM phosphate buffer. If the background is high, perform a third water wash step for 10 minutes.
• Poor water quality. Use ultrapure water (>18 megohm/cm) for washing and preparing phosphate buffer.
• Protein overloaded. Decrease the protein concentration or lower the sample volume.
• Dirty imaging platform. Always clean the imaging system with a paper towel prior to imaging the gel to minimize any background fluorescence
• Non-specific bands. Highly basic proteins and divalent metal binding proteins such as carbonic anhydrase (30 kDa), SlyD (21 kDa), and phosphorylase B (97 kDa) may cross-react with the stain producing non-specific bands.

This is likely due to overexposure - performing a longer exposure to detect low expression levels of the desired protein may result in staining of minor contaminants in the BenchMark™ His-tagged Protein Standard. Load less BenchMark™ His-tagged protein Standard or perform a short exposure to visualize and image the standard and then perform a longer exposure to visualize and image proteins expressed at low levels.

Here are possible causes and solutions:

• Improper labeling. Make sure that the labeling protocol is correctly followed to obtain the best results. Make sure you have added the Lumio™ Green Detection Reagent to the samples prior to electrophoresis. Limit exposure of the Lumio™ Gel Sample Buffer (4X) to air. Always return the Lumio™ Green Reagent and Lumio™ Enhancer to −20 degrees C immediately after use to preserve the activity of buffers.
• Low protein load or low expression level. Check total protein loaded on the gel by staining the gel with a total protein stain as described in the manual. Load at least 1 pmole of the Lumio™ fusion protein. Make sure the Lumio™ tag is in frame and the protein is expressed properly. A positive control is supplied with the Lumio™ vectors to verify the expression protocol.
• The gel is exposed to UV light for a long time. The fluorescent dye of the Lumio™ Green Reagent is sensitive to photobleaching, so avoid exposing the gel to UV light for a long time.
• The gel is not visualized immediately or imaged properly. Be sure to visualize the gel after removing the gel from the cassette and view the gel immediately after electrophoresis. Use a UV transilluminator or a laser-based scanner using appropriate filters as described in the manual.
  
  Tip: If you have run BenchMark™ Fluorescent Protein Standard on the same gel and can view the standard bands on the gel, then you are imaging the gel properly.

Here are possible causes and solutions:

• Improper handling of gels or dirty imaging platform. Avoid touching the gel with bare hands while handling or imaging the gel. Always clean the imaging platform with a paper towel prior to imaging the gel to minimize any background fluorescence.
• Protein was overloaded. Decrease the protein concentration or lower the sample volume.
• Non-specific bands. Use the Lumio™ In-Gel Detection Enhancer to minimize non-specific binding. Certain proteins from E. coli lysates (SlyD, 21 kDa) and serum proteins (BSA, 66 kDa) from the mammalian cell culture medium may cross-react with the Lumio™ Green Reagent producing non-specific bands. Removing the cell culture medium and washing the mammalian cells 3–4 times with PBS after harvesting the cells minimizes the non-specific binding from BSA.

Here are possible causes and solutions:

Here are possible causes and solutions:

This is likely due to weak cross-reaction staining of proteins containing histidine clusters. Here are our recommendations:

• Wash gel for additional time in water (step 5.)
• Slightly decrease staining time (step 2.)
• Adjust exposure time and other settings to minimize weak, non-specific staining.

For Pro-Q™ Diamond Phosphoprotein Gel Stain to work properly, it is necessary to delipidate and desalt the sample prior to electrophoresis by following the chloroform/methanol precipitation procedure in the protocol. The Pro-Q™ Diamond dye will also bind phospholipids and the dye charge interaction with phosphates can be masked by the presence of counter ions and a high salt concentration.

All SDS and fixative must be removed from the gel for optimal staining specificity. The fixation step removes the SDS and the water washes remove the fixative. To make sure that all the SDS and fixative are removed, it is necessary to do multiple changes in fixative solution followed by multiple changes in water. Larger or thicker gels may require increased volumes or incubation times in the fixative and water wash solutions, or the microwave staining procedure can be performed.

The gel may need a longer time in destain solution. Return the gel to the destain solution and continue to incubate in destain solution until only two bands are visible in the PeppermintStick™ standard lane.

This indicates that the Pro-Q™ Diamond dye has degraded and the staining solution should be discarded. Either the stain has been used past the stability period or it has been exposed to excessive room light during storage. Exposure to room light will gradually degrade the dye molecule, cleaving the phosphate-binding moiety and turning the dye into a non-specific protein stain. This will happen before the dye photobleaches, although the overall signal should be weaker than the specific signal obtained with non-degraded dye. It is likely not possible to save the stained gel, but you could try completely removing the dye by repeating the fixation step overnight, washing in water to remove fixative and then re-staining using a new stock of Pro-Q™ Diamond Phosphoprotein Gel Stain.

Many total protein stains including SYPRO™ Ruby Gel Stain and Coomassie™ Blue stain will quench the Pro-Q™ Diamond signal. If you are staining your gels or blots with Pro-Q™ Diamond stain in containers that have previously been used for a total protein stain, you may be contaminating your gel with residue left on the staining dish from the total protein stain. Either use new containers, such as plastic weigh boats, designated containers for each stain, or rinse the container well in ethanol and wipe out any residual residue with a Kimwipe™ tissue.

Over-oxidation with periodate during the step 2.5 Oxidizing Solution incubation will cause strong non-specific staining of non-glycosylated proteins. Do not incubate standard mini-gels or blots longer than 30 min, or large format 2D gels longer than one hour or use more than the recommended concentration of periodate.

If the CandyCane™ standard and test glycoproteins are staining correctly, then the kit is performing well. Some very highly abundant proteins, such as albumin in serum and plasma, may stain lightly. The Pro-Q™ Emerald dye is covalently attached to sugar residues, so more post-staining washes will help to remove any non-covalently bound dye. Non-specific staining due to high abundance can be determined by post-staining with a total protein stain, such as SYPRO™ Ruby Protein Gel Stain. That being said, some proteins are actually heavily oxidized in the native state, and this carbonylation will be picked up by the Pro-Q™ Emerald reagent. Carbonylation of amino acids can be distinguished from glycosylation by performing the Pro-Q™ Emerald staining without the oxidation step. Under these conditions, the CandyCane™ marker bands will not be stained, but the carbonylated proteins will.

Poor staining can be caused by the presence of primary amines, such as from Tris or glycine, that will also react with the aldehyde/ketone groups generated by periodate oxidation. This effectively caps the reactive groups, leaving no reactive sites for Pro-Q™ Emerald dye to bind. To remove any amine contamination, increase the volume or number of incubations in fresh fixative and then increase the volume or number of washes in wash buffer.

Poor staining can also be due to inadequate removal of the periodic acid oxidation solution. Increase the volume or number of washes after the oxidation step to ensure adequate removal of periodic acid.

Poor staining can also be due to inadequate oxidation of glycoprotein sugars. Increase the volume of periodic acid oxidation solution.

Note: Do not increase the number or incubation time of the periodic acid oxidation in excess of 30 minutes for small-format gels. Excessive periodic acid oxidation could result in increased staining of non-glycosylated proteins.

In general, large format, unusually thick or very high percent acrylamide gels may require additional incubations or wash steps for optimal signal and sensitivity of staining with Pro-Q™ Emerald Glycoprotein Gel Stain.

Many total protein stains including SYPRO™ Ruby Gel Stain and Coomassie™ Blue stain will quench the Pro-Q™ Emerald signal. If you are staining your gels or blots with Pro-Q™ Emerald stain in containers that have previously been used for a total protein stain, you may be contaminating your gel with residue left on the staining dish from the total protein stain. Either use new containers, such as plastic weigh boats, designated containers for each stain, or rinse the container well in ethanol and wipe out any residual residue with a Kimwipe™ tissue.

Speckling of Pro-Q™ Emerald dye, especially with Pro-Q™ Emerald 300 stain, can occur as the Pro-Q™ Emerald dye ages, due to self-aggregation of the dye over time. Speckles can also form due to dye binding to contaminants from the staining container, solutions, or particles from the air or gloves. Non-dye speckles can also show up in the image from auto-fluorescent particles of dust, hair, glove powder, or clothing lint that falls on the gel or surface of the glass imaging plate. The better the imager is at focusing on surface features of the gel, the more speckles that are going to be visible.

To minimize the formation of speckles and other background debris, follow clean laboratory practices, use ultrapure water of >18 megohm-cm resistance to prepare solutions, rinse gloves in water to remove powder residue before touching gels, use lint-free wipes and wear a lab coat or avoid wearing clothing that generates a lot of lint, always rinse the staining container with ethanol and wipe out any residual dye before staining another gel, and always rinse and wipe down the glass imaging surface with ethanol and water before placing your gel down. Once speckles have been deposited on the gel, it is not possible to wash them off. When analyzing amounts of glycoprotein near the limit of detection, we advise running samples in the middle lanes of the gel.

Speckles will show up as sharp, tall spikes on 3D renditions of gel images. These spikes look distinct from 3D renditions of protein spots or bands. Some image analysis software packages have de-speckling algorithms that can easily identify and remove this type of pixelated noise.

Pro-Q™ Emerald Glycoprotein Gel Stain may show high background staining in Novex™ NuPAGE™ Bis-Tris and Tris-Acetate gels, especially in combination with MES running buffer or in gels that are nearing their expiration date. The gel background increases with acrylamide density and gradient gels will show a gradual increase in background from the top to the bottom of the gel corresponding to the acrylamide gradient. Increasing the number of washes or modifying incubation times will not help to reduce this background. Better results will be obtained with Tris-Glycine or Tricine gels. If you wish to continue using Pro-Q™ Emerald stain with Novex™ NuPAGE™ Bis-Tris gels, we recommend using recently purchased gels and MOPS running buffer. Glycoproteins will still be detected in gels with high background, but with reduced sensitivity.

Here are possible causes and solutions:

This is likely due to the stain time being too long. We recommend decreasing the stain time by 30 sec or 1 min.

Here are possible causes and solutions: