Invitrogen™ Novex™ Tris-Glycine Plus Midi Protein Gels, 4 to 12%, 1.0 mm

Yes, the current Tris-Glycine Midi Gels will be discontinued on December 31 2017.

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

It may be possible to reduce background by using the protocol provided for NuPAGE Invitrogen Bis-Tris gels/Small Peptides. This protocol incorporates an extra fix step to remove excess SDS, which can act as an anti-colloidal agent and lead to higher background. The low pH of the staining solution will fix the gel, but not as fast as the pre-fix step specified in the NuPAGE protocol.

*Double check that the tape on the bottom of the gel has been removed.
*Make sure that the gel(s) are oriented so that the taller sides of the cassette (with the printing) are facing the outside of the electrophoresis unit.
*Make sure that the inner buffer chamber is filled sufficiently so that the wells are covered with buffer. If the wells are not covered, check for leaks and reseal.
*Double check to see if there are any loose electrodes or connections on the Mini cell unit.
*Check the power supply unit.

You may purchase the ZOOM adapters, Cat. No. ZA10001 to help you connect your leads to the power supply

We recommend marking the cassette at the bottom of the wells with a marker pen prior to assembling the Upper buffer chamber. Also, we recommend illuminating the bench area with a light source placed directly behind the XCell SureLock unit.

Here are possible causes and solutions:

1) Buffers are too dilute: Check buffer recipe; remake if necessary.
2) Upper buffer chamber is leaking: Make sure the buffer core is firmly seated, the gaskets are in place and the gel tension lever is locked.
3) Voltage is set too low: Set correct volatage

This can happen if Tris-Glycine gels are run using NuPAGE Running buffer containing Antioxidant. Please make sure that the correct Tris-Glycine Running buffer is used with Tris-Glycine gels.

This could be due to:

*Debris in the well
*High salt in the sample (make sure that the salt concentration does not exceed 50-100 mM)
*Running buffer issue
*Gel casting error

This could be due to a gel polymerization issue combined with incorrect sample preparation (final sample dilution less than 1X). Please try a different lot of the same gel and make sure that the sample is correctly prepared.

Possible cause:

*Excess reducing agent (beta-mercaptoethanol)
*Skin protein contaminants (keratin)

Remedy:

*The addition of iodoacetamide to the equilibration buffer just before applying the sample to the gel has been shown to eliminate these artifact bands.
*Use new electrophoretic solutions and wear gloves when handling and loading the gel. This issue is more common when highly sensitive stains are used.

Possible cause:

*Carry-over contamination of sample from one well into neighboring wells due to loading error
*Contaminated running buffer
*Gel casting error: malformed wells

Remedy:

*Use a gel loading tip to load wells
*Reduce the sample volume
*Do not delay while loading wells
*Do not delay after the run, as proteins can diffuse horizontally; a full well left next to an empty well would eventually contaminate the empty well over time.

Possible cause:

*Poor polymerization around sample wells
*High salt concentration in sample
*Uneven gel interface
*Excessive pressure applied to the gel plates when the gel is placed into the clamp assembly
*Uneven heating of the gel
*Insoluble material in the gel or inconsistent pore size throughout the gel
*Air bubble during the run

Remedy:

*Remove excess salt/other material by dialysis, Sephadex G-25 or any other desalting column or using an Amicon concentrator.
*Either use a cooled apparatus or reduce the current at which electrophoresis is performed.

Gel lifting off the cassette can be caused by:

*Expired gels that are degrading
*Improper storage of gels
*Too much heat accumulating during the electrophoresis run due to excessive current
*Insufficient polymerization of the polyacrylamide

Ghost bands are usually attributed to a slight lifting of the gel from the cassette, which results in the trickling down of some sample beyond its normal migration point. It then accumulates and appears as a faint second band.

"Smiling" bands may be the result of the acrylamide in the gel breaking down, leaving less of a matrix for the proteins to migrate. We recommend checking to ensure that the gels have not been used past their expiration date.

Barbell shaped bands are a result of loading too large of a sample volume. When a large sample volume is loaded, part of the sample tends to diffuse to the sides of the wells. When the run begins and the sample moves through the stacking portion of the gel, the sample will incompletely stack causing a slight retardation of the portion of the sample that diffused to the sides of the wells. This effect may be intensified for larger proteins, whose migration is more impeded in the low concentration acrylamide of the stacking gel. To alleviate the problem, we recommend concentrating the protein and loading a smaller volume. This gives a "thinner" starting zone.

Here are possible causes and solutions:

1) Sample overload: Do not overload samples
2) Addition of reducing agent that is not fresh: Reduce samples right before loading and do not use samples that have been stored in reducing agent
3)Re-oxidation of the protein during the run: Add antioxidant to the running buffer if you are running NuPAGE gels
4) Presence of highly hydrophobic regions where the protein can exclude SDS: Load the sample with 2X sample buffer instead of 1X sample buffer
5) Excess salt in the sample: Precipitate and reconstitute in lower salt buffer
6) Not enough SDS in the sample: Add SDS to the upper buffer chamber (try 0.1%, 0.2%, 0.3% and 0.4% SDS)