SDS-PAGE gels and western blotting

9/23/04

 

2. SDS-PAGE gels

 

  1. Make sure all samples are prepared in SDS sample buffer.

 

  1. Make rainbow MW markers (Amersham) in SB also. About 5-10uL standards for one lane.

 

  1. Plan out the order of the lanes and write them down.

 

  1. Wash glass gel plates with soap and distilled water, rinse in water, and rinse in 70% EtOH. Dry with paper towels and leave standing to dry fully.

 

  1. Assemble one large and one small glass plate with 2 thick spacers in between. Place in one plate holder with the small plate on the open side. When plates are flush on a hard flat surface, screw in place.

 

  1. Clamp plates and holder in gel caster. Make sure caster has both orange and gray cushions, and a folded piece of parafilm. You may want to check for leaks by pipetting in a few mL of water. If it does not leak, pour water out into a paper towel held over the plates to absorb as much as possible.

 

  1. Mix gel according to the recipes here (see below). Chose the gel percentage based on the size of the proteins you want to see. For small proteins use a higher percentage. 15kD is good on 12-15% gels, 45kD on 10-12% gels, 96kD on 8%, 200kD consider 6%, running the gel really long, or a 4-20% gradient gel.

 

  1. For a thick gel you need 10mL of the separating gel and 4mL of the stack. Mix water, buffer and acrylamide. Only just before you are ready to pour add (for each 10mL gel) 100uL APS and 10uL TEMED. This will cause the gel to polymerize so you only have a few minutes to pour after adding. Mix by inverting the tube serveral times. Use a 10mL pipette to add gel up to about 3/4 of an inch from the top of the plates.

 

  1. Add 1ml water-saturated iso-butanol (use the top phase) over the gel. This will help it set evenly without bubbles.

 

  1. Once the gel has set (10 min or so), pour off the iso-butanol into a paper towel held over the plates.

 

  1. Now add APS and TEMED to the stack gel, invert the tube, and add to plates filling to the top. Put comb in. Make sure stack gel comes to the top of the comb. Allow to set.

 

  1. Make up 1X Tris-glycine SDS running buffer. You need 800-1000 mL/ gel. Clamp plate holders into central holder with the small plate facing in. You must have a plate holder on each side even if you are only running one gel. Pull combs and check integrity of well. Place in box and add buffer up to the top of the plate holders.

 

  1. Boil samples 2-3Õ. Cool and spin 14K 3Õ.

 

  1. Load samples into wells with gel loading tips. For a 10 well comb and a thick gel you can get about 40uL in a well.

 

  1. Run the gel at 100-200V until the dye front reaches the bottom (20-60 minutes depending on voltage and gel percentage).

 

  1. If you do not want to western, but only want to see the proteins, move gel to Coomassie stain, microwave 20 seconds, stain up to 1 additional hour, then pour off Coomassie. Destain gel with destain and knotted Kimwipes for several hours to O/N on shaker.

 

  1. If you want to western, switch to that protocol.

 

 

Recipes:

 

APS (ammonium persulfate)

1g in 10mL ddH2O

Store at 4 for 1-2 months. If your gels are setting more slowly than usual, replace.

 

1.5M Tris with 0.4%SDS

91g Tris base in 300mL ddH2O

pH to 8.8 with about 12.75mL 12.5N HCl

dd to 500mL

filter, add 2g SDS

store at 4 for up to 3 months

 

0.5M Tris with 0.4%SDS

6.05g Tris base in 40mL ddH2O

pH to 6.8 with about 3.5 mL 12.5N HCl

dd to 100mL

filter, add 0.4g SDS

stroe at 4 for up to 3 months

 

Separating gel

 

8%

10%

12%

15%

ddH2O

4.7mL

4.1mL

3.4mL

2.4mL

1.5M Tris 8.8

2.5mL

2.5mL

2.5mL

2.5mL

30% acrylamide

2.7mL

3.3mL

4.0mL

5.0mL

 

Stacking gel

3.5mL ddH2O

1.5mL 0.5M Tris 6.8
1mL acrylamide

 

2X SDS-sample buffer

2.5mL 0.5M Tris with 0.4% SDS

2mL 10% SDS

2.5mL 80% glycerol

1mL B-mercaptoethanol

2mL ddH2O

Bromphenol Blue to color