Version 1/96

Silver-Stained Laemmli Gels: How to Avoid High Background

During the last stages of purification of an enzyme, you may need to silver stain the SDS-PAGE Laemmli gel for the analysis of your column fractions. If you don't take special precautions you will almost always have background problems, most notably are bands in the ‰ 60 kDa region of the gel which may be keratin. After many trials, Li Liu in the Gelb lab has come up with the following protocols which seem to reproducibly give beautiful silver-stained gels with low background.

Preparing the Laemmli Gel:

1. Before the column that will generate the fractions to be analyzed by silver-stained SDS-PAGE, handle the fraction collector tubes with gloves (Baxter # 8856). All buffers used to prequilibrate and run the last column should be filtered using the glass, 2-piece filter device used for filtering FPLC and HPLC solvents. The apparatus should be rinsed well with MQ-water and a new filter disk should be used (Micro Separations Inc. # N04SP04700 Nylon 0.45 u). All of these steps (assembling and washing the filtration device, the filter flask receiving the filtered buffers, and the buffer storage vessels) should be done with gloves.

2. TCA/deoxycholate is often used to precipitate small amounts of protein prior to SDS-PAGE. Samples prepared in this way usually have a high background when the gel is silver stained. Thus if you want low background you cannot use this procedure; it is better to take aliquots from the column fraction tubes and treat them directly with Laemmli sample buffer. You may be forced to load a larger than normal volume of sample onto the gel lanes.

3. CRITICAL: Laemmli sample buffer must be made with reagents than have not been opened before. To guarantee this, you should order new bottles of reagents. In this case make a large volume of sample buffer, perhaps 50 ml, so that this can be stored in aliquots for others in the lab to use. Each person should have their own aliquot of sample buffer so that they can guarantee that it will not become contaminated. You should use the following reagents: SDS (ICN # 811036), MQ water, glycerol (Fisher Biotech # BP229-1), bromophenol blue (BioRad # 161-0404), beta-mercaptoethanol (Fisher Biotech # BP176-100) or DTT (BioRad # 161-0611. After mixing all components, filter the solution with a new syringe filter cartridge (Millex-GS) hooked up to a new disposable plastic syringe into new Falcon tubes. Store the closed tubes at -20°C (they have been stored in this way for 1 year without problems). Filter a small amount of sample buffer as above just before use. Use gloves for all the steps!

4. Use the following reagents for the various Laemmli buffers: glycine (USB # 16405), acrylamide/bis-acrylamide mix (Boehringer Mannheim # 11685 821), Tris (USB # 22674). You don't have to use new bottles of these reagents. Glycine and Tris are stored at room temperature and the acrylamide/bis-acrylamide stock solution is stored at 4°C. Tank buffer is made and then filtered using the 2-piece, HPLC/FPLC solvent filter device with a new membrane (see step 1 above). Tank buffer is stored at room temperature in a plastic bottle that was previously rinsed well with MQ water and autoclaved. Keep your own private stocks of tank buffer, again to guarantee against contamination. Each time you run a new gel, freshly filter the tank buffer. Wear gloves during all steps!

5. Running gel and stacking gel buffers are made as in step 4 and are stored in Pyrex tissue culture bottles (the ones with the orange plastic screw caps from VWR). Before use, rinse the bottle well with MQ water and autoclave. They can be stored as in step 4 above at room temperature. Keep your own private stocks of buffers, again to guarantee against contamination. Wear gloves during all steps!

6. The gel plates, spacers, electrode assembly, and comb are washed with Liqui-Nox detergent (VWR), then washed well with MQ water. The plates are placed on a KimWipe and dried in the ‰ 50°C oven, the spacers and comb are wiped dry with a KimWipe. Wear gloves during all steps!

7. Before pouring your gel, make up solutions of stacking gel and running gel, and use these immediately to pour the gel, i.e. the buffers used to make these solutions can be stored at room temperature as indicated in steps 4 and 5 above, but after you mix together all the ingredients (buffer, acrylamide/bis-acrylamide stock, ammonium persulfate, TEMED) you should not store this mixture. After mixing the components, filter the mixtures via syringe-cartridge and syringe as in step 3 above, and use the filtered solutions to pour the gel. Ammonium sulfate is BioRad # 161-0700 and TEMED is BioRad # 161-0800. Wear gloves during all steps!

8. The eppendorf tubes that you use for boiling sample with sample buffer should be previously autoclaved. Load the samples onto the gel with the long Pipettor tips (Perfect Scientific # 1590). These tips are loaded into a plastic rack and autoclaved. Wear gloves during all steps!

Silver Staining The Gel:

Based on: T. Rabilloud et al., Electrophoresis 9, 288, 1988.

By trial and error, the Gelb lab has found that the following protocol is sensitive and has a low background if followed carefully. Read the entire protocol before starting! The gel is placed in a plastic box that can be closed with a lid (i.e. Tupper-Ware) and 50 ml of each solution for a mini-gel or 200 ml of each solution for a 15 x 15 cm gel are used. When you change to a solution with a new composition, pour a bit of the new solution over the gel and briefly tilt the container so that the side walls get rinsed, then drain, then proceed as indicated below for each step (thus, for each step below there is actually n + 1 rinses, where n is the number of rinses indicated below). All solutions are made with milli-Q water. If you have to handle the gel, wear gloves!

1. Fix the gel in 10% acetic acid, 30% ethanol (pint size bottle of absolute ethanol) overnight (or 4 hr).

2. Rinse with 30% ethanol (3 x 20 min) and with water (2 x 15 min).

3. Sensitize the gel with a solution of 0.25 g/liter sodium dithionite (Na2S2O4) for 1 min (make this solution fresh during the end of step 2 and use immediately). Rinse with water (2 x 1 min). Prolonged reaction times in Na2S2O4 produces higher background staining.

4. Incubate the gel for 20-30 min in 0.2% AgNO3 solution containing 80 ul/liter 37% formaldehyde (Aldrich 25, 254-9, avoid using an old bottle of yellowed formaldehyde) (make this fresh during step 2). Rinse with water for 1 min.

5. Develop 2 x 5 min in 4% K2CO3 solution containing 500 ul/liter 37% formaldehyde and 5 mg/liter sodium thiosulfate (Na2S2O3) (this solution can be made in advanced and stored at RT for < 2 weeks). Watch the gel carefully, protein bands should appear within few mins, prolonged incubation leads to high background staining.

6. To stop the developing process, drain off developing solution and add solution of a new portion of 100 ml of developing solution containing 3.5 ml acetic acid.

7. Rinse gel with water (4 x 30 min).

8. For your lab notebook you should photograph the gel shortly after drying it down because the gel will darken slowly over time.