Lidstrom: SDS-PAGE

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Gel Prep

  • Clean cover plate and thicker spacer plate 75 mM
    • Soap and Water
    • Ethanol
    • DI water
  • Dry plates
  • Setup one spacer plate and one cover plate in each gel holder
    • The cover plate goes on the side of the spacer plate with the spacers in order to create a small gap between the plates
  • Put the gel holder into the casting stand

Pour the Resolving Gel

  • Mix components of the amounts in the Gel Mix link for the resolving gel (Recipes is for 4 gels). Mix in the order listed.
  • Gel Mix Recipe
    • Don't add APS/TEMED until ready to pour
    • The APS solution should be ~ 1-2 months old. A 3 month old solution failed to cause polymerization -JM 10/2012
  • Use pipette to put gel mix into the gap between the plates
  • Carefully layer 50%EtOH 50% ddH2O on top of the gel to prevent the top of the gel from drying out
  • Let dry for an hour
  • Store at 4 deg C wrapped in a wet paper towel and saran wrap if you're not going to use it right away.

Pour the Stacking Gel

  • Replace gel in gel holder
  • Dry surface of gel carefully with Kimwipe or paper towel
    • It can be a little gooey
  • Mix components of the amounts in the Gel Mix link. Mix in the order listed.
  • Gel Mix Recipe
    • Don't add APS/TEMED until ready to pour
  • Use pipette to put gel mix into the gap between the plates
  • Insert the comb being 'careful not to trap any bubbles'
  • Attach binder clips to help hold the comb in while drying. One in either side of the casting stand clamp.
  • Leave for 1 hour while polymerization occurs.
  • Can store for a few weeks in the fridge. Leave comb in, and wrap in a wet paper towel and cling wrap.
binder clips squeeze the glass to the comb. Put them as far down as they go.

Sample Prep

  • 10-20 ug protein
  • Load 12-15 uL, absolute max 20 uL for big comb
  • Denature protein
    • Mix with sample buffer & BME
      • How much?
    • Boil for 5-10 min (can do longer times at lower temp, too.)

Running the Gel

  • Bio-Rad Mini-Cell Setup
    • If only 1 gel, use buffer dam to replace second gel
  • Slot gels with cover plates facing each other...
  • Apply pressure on gel holder and gels as you close the tabs to seal the center compartment.
Mini-cell Gel holder
  • Fill central compartment with running buffer
  • Pour rest into outer compartment
  • Load gel
  • Make sure to color/charge-match the cords to the power unit as the electrodes in the gel holder to the contacts in the lid.
  • Run 30-45 min @250V
    • Amanda runs 20 min at 200V, then checks frequently to make sure you don't run your protein off the gel.

Staining, Destaining, & Visualization

  • dye overnight or cycles of 1 min @ power 6 in the microwave
    • microwave by Bo's bench. Let it vent a little in the hood between heating events.
  • Return dye to container
  • Rinse to remove residual dye
  • Destain (I do 2 rounds at least)

Other Resources

Mistakes to Be Careful About

  • letting the gel dry too long after pouring the stacking gel (comb step)
    • the very edges can shrivel up, which becomes a problem when you try to use those edge lanes
  • sample sloshing out of the well you are using into a neighboring well

Recipes

All recipes except the staining & destaining solution are from the Mini-PROTEAN® Tetra Cell manual

Loading Buffer (5x):

    • You will mix pre-made 5X loading buffer with fresh beta-mercaptoethanol prior to each use.
    • 5x loading buffer: 3.55 mL deionized water, 1.25 mL 0.5 M Tris-HCl pH 6.8, 2.5 mL glycerol, 2.0 mL of 10% (w/v) SDS, 0.2 mL of 0.5% (w/v) Bromophenol Blue. Total volume = 9.5 mL.
  • mix 50 uL beta-mercaptoethanol to 950 uL sample buffer prior to use.
    • Scaled back 10x: 10 uL beta-mercaptoethanol + 190 uL 5x buffer
  • Dilute the sample at least 1:2 with sample buffer and heat at 95oC for 4 min to lyse the cells.
  • Amanda mixes 160 uL 5X buffer, 8 uL beta-mercapto-ethanol, 32 uL H2O. Use 10 uL per 25 uL of cells.
  • ?? Should add up to 8M urea for really hydrophobic proteins

Running Buffer:

  • 10x SDS running buffer (1 liter): 30.3 g Tris-HCl, 144 g Glycine, 10 g SDS. Fill to 1 L with ddH2O. Don't add acid or base to adjust pH.
  • Make 1L of 1x for use.
  • Store at 4oC.
  • This buffer is used while running proteins through the gel. Pour it in as the instructions for the box explain. Pour back into bottle for re-use afterward.
  • Keep until ____; remake after this.

Staining Buffer(Coomassie Brilliant Blue G-250):

  • 0.500 g Brilliant Blue, 500 mL methanol, 100 mL glacial acetic acid, 400 mL dH20. Mix well. Store at room temperature; can be reused 2-3 times.
  • Amanda's recipe: 0.4 g of Coomassie Blue R 350 in 200 mL of 40% (v/v) methanol in water. Stir & filter (coffee filter is fine). Add 200 mL of 20% acetic acid in water (40 mL acetic acid in 160 mL water).
  • note: Coomassie Brilliant Blue G-250 differs from Coomassie Brilliant Blue R-250 by the addition of two methyl groups. We use the G form. Read more about the R form here or at the bottom of this page.

Destaining Buffer:

  • 30% methanol, 10% acetic acid, water

Acrylamide toxicity

  • Acrylamide is toxic to your nervous system, and may be a carcinogen. The unpolymerized form is toxic, but the polymerized form is much less toxic. ALWAYS wear gloves and wipe up spills - once the solution drys, the dust can be inhaled. Interestingly, fried starchy/sugary foods naturally contain acrylamide, too.
  • more than you want to know about acrylamide toxicity can be found here

Ladder

PageRuler protein gel legend

The two types of Coomassie Blue dyes

    • note: Coomassie Brilliant Blue G-250 differs from Coomassie Brilliant Blue R-250 by the addition of two methyl groups. We use the G form. Read more about the R form here.
  • R in R-250 stands for Reddish hue while G in G-250 for Greenish hue. R-250 is dark reddish blue/purple stain while G-250 gives lighter greenish blue stain.

BioRad once told Amanda:

  • The G-250 form the colloidal particles in an aqueous solution. This is an advantage for staining a gel because the colloids tend not to stain the gel matrices, reducing the background problem. When the colloids come close to the proteins, the dye molecule is removed from the colloids by the nearby proteins due to the higher affinity of proteins to the dye.
  • R-250, on the other hand, doesn't form the colloids. Rather, an individual dye molecule is dispersed in a solution. Therefore, the dye molecules can interact not only with proteins but with gel matrices freely, creating the background staining issue.
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