Nucleic Acid Gel Electrophoresis and Blotting Support—Troubleshooting

First check the percentage of your agarose gel. A higher percentage will help you to resolve smaller molecular weights while a lower percentage will help you to resolve larger molecular weights.

Check the power supply to ensure current is being passed through the gel. The running buffer composition must match the gel chemistry.

Usually, fuzzy bands are caused by incorrect buffer solutions. Check the pH and salt concentrations in the running and gel buffers. Fuzzy bands can also be caused by excess salt in the samples or by old gels.

We would recommend running the gel for longer or choosing a different gel that will have higher resolution in the size range you are targeting.

Do not run the gels longer than 60 minutes for the single comb gels or 25 minutes for the double comb gels. Ions are depleted and longer run times will damage the gel. Do not run E-Gel® 48 Agarose Gels longer than 30 minutes or E-Gel® 96 Agarose Gels longer than 20 minutes.

The conformation of plasmid DNA will affect its mobility on a gel. Plasmid DNA can be supercoiled (native 3D conformation), nicked circular (nick in one DNA strand) or linear (nicks in both DNA strands). You may see all three forms on an agarose gel. Supercoiled DNA runs the fastest and will show up lowest on the gel, linear DNA runs in the middle of the gel, and nicked circular DNA shows up at the top of the gel, since it migrates the slowest.

While we recommend storage at room temperature, these gels will still be usable. Bring the gels to room temperature prior to the run for optimal conditions.

Potential issues are:

• Loading too much DNA. We recommend using 20–100 ng per band of DNA; however, you can use up to a maximum of 500 ng.
• Samples with a high salt concentration. Samples containing >50 mM NaCl, 100 mM KCl, 10 mM acetate ions, or 100 mM EDTA will cause loss of resolution.
• Samples may have diluted in TAE instead of TE or water.
• Running E-Gel® Agarose Gels too long. We recommend that you run the gels for 25–30 minutes for straighter patterns. Do not run longer than 60 minutes or the gel will be damaged.
• A voltage or current that is too high. If using an E-Gel® base with power supply, run the gel at 60–70 V (constant voltage) or 40–50 mA (constant current). Do not allow the current to exceed 60 mA.
• For E-Gel® 96 Agorse Gels being run on an E-Base™ device, be sure you are running on the “EG” program rather than the “EP” program designed for E-PAGE™ gels.
• Sample that was not properly loaded or had a very low volume of sample loaded.
• The gel was not electrophoresed immediately after sample loading (for best results, run gel within 15 minutes of loading).

Please ensure that you have not overloaded the well and that the wells were not damaged during comb removal.

Check the E-Gel® cassette and copper contacts. You can try replacing the gel cassette with a fresh gel cassette to see whether this fixes the problem. A cold cassette or improper operating conditions can also lead to this failure. Cassettes should be at room temperature for use; avoid storing at 4°C.

Here are some suggestions:

• Try cleaning the cassettes with alcohol and Kimwipes® wipers.
• Try cleaning the camera lens.
• Try to adjust the exposure time and brightness options of the documentation system you are using.

Please check the cord you are using for the iBase™ Power System. Oftentimes, the power cords for the PowerBase™ v.4 system and the E-Gel® Safe Imager™ Real-Time Transilluminator get mixed up. If this is the case, the blue light will not come on even though the fan and LED light will operate. You will need to use the iBase™ cord, which should say 48 V on it.

Here are some common reasons why your gel is not running properly:

• Copper contacts in the base are damaged due to improper use. Make sure the copper contacts in the base are intact.
• An expired or defective gel cassette was used.
• The E-Gel® EX cassette was not inserted properly into a base.
• An incorrect adaptor was used.

Here are some possibilities and suggestions to resolve the problem:

• The sample is overloaded. Do not load more than the recommended amount.
• A high salt concentration. Dilute the samples.
• The E-Gel® EX Agarose Gels were prerun; this is not recommended for these gels.
• A very low volume of sample was loaded or the sample was not loaded properly.
• Bubbles may have been introduced while loading the samples. Bubbles will cause band distortion. Load the recommended sample volume based on the gel type and loading method. For proper band separation, we recommend keeping sample volumes uniform. Load deionized water or TE into any empty wells, and avoid introducing bubbles.
• The gel was not electrophoresed immediately after sample loading. Run your sample within 1 minute of loading it.
• The E-Gel® Agarose Gel may have been used beyond its expiration date. Check the expiration date.
• A longer electrophoresis run time or high current was used during the run. Longer run times cause an increase in the current, resulting in poor band migration or a melted gel. Do not run the gel longer than the recommended time for each gel type.

We do not currently offer software that is compatible with Mac® computers. Please install the software on a 32-bit or 64-bit PC running either a Windows® XP Professional or Windows® 7 Professional operating system.

This error can occur if there is other software running in the background, like an Internet Explorer® browser, or if there is an antivirus program running that is limiting his ability to access some files.
To fix this error, try to:

• Turn off Internet Explorer® or any other program running during the install.
• Inactivate the antivirus program (or  temporarily uninstall it), install the E-Gel® Imager software, then reactivate or reinstall the antivirus program.
• When the error comes up, select "Ignore" to allow the installation to continue instead of "Retry" or "Abort".

Depress the activation button for at least 2 seconds, and the imager will transilluminate for 5 minutes rather than 30 seconds.

1) Check that the camera hood or activator key is in place to make sure the base is not deactivated.
2) Light bases will "time out," but the power indicator light may stay illuminated. Turn the base off, then back on.

Follow recommended DNA dilutions and leave the gel to cool down on the bench or for a few minutes in the fridge. Please check troubleshooting tips provided in the manual.

If a slight turbidity develops, the fine precipitate can be dissolved by autoclaving for 5 minutes at 110°C. Do not autoclave in the container supplied. This treatment has no deleterious effect on the buffering properties of TBE.

Here are some suggestions for your experiments:

• The RNA samples may have been degraded by RNases. Use standard precautions to prevent contamination.
• Make sure samples are heated for 3 minutes at 70°C just prior to loading. If this is not done, the oligonucleotides will not be fully denatured, which may result in a smeared background.
• Be sure to vigorously flush urea out of sample wells just prior to loading the sample. Urea will continually seep from the gel into the well. Urea is very dense and will force the sample into a ball.
• A sample volume over 10 µL may result in smearing. This volume is less than may be used with agarose gels.
• If ultrapure water (18 milliohms) is not used, smeared bands and high background may result.
• Check to make sure the running and sample buffers have been prepared and diluted correctly.
• Make sure to run the bromophenol blue until it reaches the slot. Stopping the run short can result in less than optimal results.

You can use the Reverse E-Gel® Program to run the band back into the collection gel.

Please see below the top ten ways to increase sensitivity of your Northern hybridizations:

1) Increase the amount of RNA loaded in each lane (up to 30 mg). 2) Use poly(A) RNA instead of total RNA; 10 mg of poly(A) RNA is ~300–350 mg total RNA (3–5%). 3) Switch to ULTRAhyb® Ultrasensitive Hybridization Buffer. 4) Switch from DNA to RNA probes. 5) Use downward alkaline capillary transfer. 6) Use an optimal hybridization temperature. 7) Use a freshly synthesized probe. 8) Use a high specific activity probe (10^8 to 10^9 cpm/mg). 9) Increase exposure time (it can take up to 3 days to see low-abundance messages with radiolabeled probes). 10) Follow the manufacturer’s recommendations to crosslink the RNA to the membrane.

Read more about these suggestions here.

For RNA probes on DNA or RNA targets: Autoclave the membrane in a bottle containing 0.1% SDS solution for 15 minutes. Repeat if necessary. For DNA probes on DNA targets only: You can use the same protocol used for RNA probe stripping. Another option is alkaline denaturation. Incubate the membrane with 400 mM NaOH for 30 minutes, then wash with 0.1% SDS for 15 minutes. These stripping methods should work for 2–3 stripping procedures. However, nucleic acids will gradually be removed from the blot.

rRNA makes up ~80% of total RNA samples. When 10 µg of total RNA is loaded into a Northern gel lane, the 18S and 28S rRNA bands contain 2–6 µg RNA each. This amount of nucleic acid can nonspecifically trap probe as well as bind complementary sequence. Probe trapping by rRNA can be reduced by using the minimal amount of probe, and by labeling only sequence complementary to mRNA. Transfer using a basic buffer can prevent trapping. Finally, you can use a high hybridization and wash temperature to minimize cross hybridization to rRNA.

Residual RNA could be due to:

• An inadequate volume of transfer buffer (>0.5 mL/cm2)
• Too much weight.
• The gel used was too thick (>6 mm).
• The transfer time was insufficient (we recommend 20 minutes/mm)

Poor signal could be a result of the following:

• A hybridization temperature that was not optimal.
• Probe degradation (too old).
• A low specific activity probe (should be ~2 x 10^9 cpm/ug, random primed).
• A probe that was not denatured (DNA).
• A probe concentration that was too low (<10^6 cpm/mL).
• A longer hybridization time needed.
• Poor transfer of RNA to membrane.
• Inadequate cross-linking or overexposure to UV light.
• An alkaline transfer time that was too long (>4 hours).
• The wrong membrane (nitrocellulose).
• Failure to follow nonisotopic detection protocols.
• A message that co-migrates with ribosomal RNA.
• Inappropriate use of intensifying screens.

There are several types of background, and each can have a different cause:

1) Blotchy signal across the membrane: This can be caused by a membrane of poor quality, one that has dried out, or one that has been mishandled (e.g., oil from human skin, powder from gloves). Use high quality nylon membrane that has not previously been handled and use forceps to handle the membrane from the edges. Blotchiness can also be caused by uneven distribution of the hybridization reagents. Do not pipette probe directly onto the membrane in hybridization solution; dilute it into the hybridization solution first.

2) A smear through the lane: Hybridization conditions that are substantially below the optimum for a given probe can lead to high lane-specific background and/or substantial cross-hybridization. Start with a high hybridization temperature and slowly decrease the temperature until a specific signal is obtained. High probe concentrations, especially for nonisotopic probes, can also cause lane-specific background. Use 10 pM nonisotopically labeled DNA probes and 0.1 nM nonisotopically labeled RNA probes.

3) Speckling across the membrane: Probe preparations with poor incorporation (or where unincorporated nucleotides have not been removed) can cause speckling on the membrane. Check probe quality and remove unincorporated nucleotides. Particulates in probe preparations or hybridization buffer (e.g., when not completely in solution) can also cause speckling on the membrane. Ensure that these reagents are in solution, and consider centrifuging in a microfuge or low-speed centrifuge, or filtering the solutions through a 0.22 µm filter to remove particulates.

If you see high background that is not associated with the lanes, this could be due to:

• A bad membrane or incompatible membrane.
• A membrane that dried out during procedure.
• Reagents that were not evenly distributed.
• Microbial contamination.
• Particulate matter deposited on the membrane.
• Precipitates present in nonisotopic detection reagents.
• Agarose or transfer buffer that dried on the membrane.
• Static charges developing during film development.
• A blot that was too wet when exposed to film.

The following reasons could have led to cross-hybridization:

• The probe concentration was too high.
• Hybridization/washing conditions were not stringent enough.
• There were multiple targets in the mRNA.
• There was too much nonhomologous sequence in the probe.
• Cross-hybridization to ribosomal bands. This can occur when the total RNA has a large amount of rRNA on the blot, trapping the probe. In this case, you should see the specific band, but it may be much fainter.
• The hybridization temperature was too low (try increasing it up to 52°C).