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
• Make a fresh 10% (w/vol) APS solution. Don't use an "old" solution; the gel won't polymerize. 10% w/v = 0.1 g/mL.

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
• Rinse off ethanol/water or butanol mix used to keep the top of the gel hydrated
  • not rinsing will result in bubbles between the gel and the plate (bad)
• 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

• use 10-20 ug protein. Can protein normalize and add the same amount of total protein across cultures.
  • Obtain by boiling cultures with concentrated sample buffer + beta-mercaptoethanol
    • need to prepare this mix from concentrated loading dye (pre-made) and beta-mercaptoethanol. Make this mix fresh each day.
  • If using Amanda's mix, mix 10 uL of this mix (proportions below) into every 25 uL of cell culture.
  • If using BioRad's mix, "dilute the samples at least 1:2 with sample buffer", which I presume is 1 part cell culture + 2 parts mix.
  • Heat your sample by either: (link)
    • a) Boiling for 5-10 minutes (Works for most proteins)
    • b) 65 degrees C for 10 minutes (If you have smearing using the above procedure)
    • c) 37 degrees for 30 minutes (Membrane proteins or others that do not enter the gel otherwise may benefit from this type of sample preparation)
• Load 12-15 uL, absolute max 20 uL for big comb
  • less for the skinny wells
• If gel is overloaded or underloaded, run a new gel with a different amount of dyed & boiled lysate

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
  • should fill sample wells
• Pour rest into outer compartment
  • fill to specified line
• 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 @ 60 V for __ min, then 200 V for ~ 20+ min.
  • Can skip the 60V step if you don't need a gorgeous gel
  • 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

• Background (Wikipedia)
• Instructions for PageRuler protein standard
• Instructions for BioRad Rainbow Std
• Mini-PROTEAN® Tetra Cell manual

Mistakes to Be Careful About

• using an "old" APS solution when making gels. The 10% weight/volume APS should be made fresh each time for best results. Don't use a solution that is more than a month or so old. The gels won't polymerize.
• not mixing the liquid gel mixture enough
• 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
• using too much beta-mercaptoethanol in your sammple buffer
  • should have < 1% beta-mercaptoethanol in the mix after you add sample buffer to the
  • too much reduction of cysteines is bad: will alter structure and even cleave proteins.

Recipes

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

Loading Buffer

• You will mix pre-made concentrated loading buffer with fresh beta-mercaptoethanol prior to each use.
  • BioRad's loading buffer recipe: 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 5x: 10 uL beta-mercaptoethanol + 190 uL 5x buffer
  • scaled back 10x: 5 uL beta-mercaptoethanol + 95 uL buffer
• Dilute the sample "at least 1:2 with sample buffer" and heat at 95oC for 4 min to lyse the cells.
  • Janet presumes this is 1 part sample per 2 parts buffer, but isn't sure (???)
• Amanda's recipe is a little different: 4x mix made of: 4 mL glycerol, 0.8 g SDS, 2.5 mL 1M Tris-HCl pH 6.8, 80 uL of bromophenol blue slurry (5 mg/mL = 0.5% (w/v)), H2O to 8 mL.
  • Amanda mixes 160 uL 5X buffer, 8 uL beta-mercapto-ethanol, 32 uL H2O. Use 10 uL per 25 uL of cells. Recipe Comparison (to BioRad's)
• Can 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):

• recipes vary. Usually there is about 0.5 g/L dye, and between 200 - 500 mL methanol per liter. 100 mL/L acetic acid is very common.
• 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
• some labs use much less methanol & acetic acid; some use plain water.
• Janet rinses in plain water before using our destianing buffer.

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.