Lidstrom: SDS-PAGE: Difference between revisions
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**To make this buffer, add: | **To make this buffer, add: | ||
***blah | ***blah | ||
* Running Buffer: | * Running Buffer: | ||
** 10x SDS running buffer (1 liter): 30.2 g Tris-HCl, 144 g Glycine, 10 g SDS, adjust to pH to 8.3; bring volume up to 1L with dH20; store at room temperature. Dilute to 1x with dH20. | |||
** 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. | ** 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. | ** Keep until ____; remake after this. | ||
* Staining Buffer: | |||
** | * 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. | |||
** 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 [http://en.wikipedia.org/wiki/Coomassie_Brilliant_Blue here]. | ** 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 [http://en.wikipedia.org/wiki/Coomassie_Brilliant_Blue here]. | ||
* Destaining Buffer: | * Destaining Buffer: | ||
* | * |
Revision as of 13:11, 22 October 2012
Return: Protocols
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.
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.)
- Mix with sample buffer & BME
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.
- 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
- Background (Wikipedia)
- Instructions for PageRuler protein standard
- Instructions for BioRad Rainbow Std
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
- Loading Buffer (5x):
- 10% w/v SDS, 10 mM Dithiothreitol or beta-mercapto-ethanol, 20 % v/v Glycerol, 0.2 M Tris-HCl, pH 6.8, 0.05% w/v Bromophenolblue
- Should add up to 8M urea for really hydrophobic proteins
- To make this buffer, add:
- blah
- Running Buffer:
- 10x SDS running buffer (1 liter): 30.2 g Tris-HCl, 144 g Glycine, 10 g SDS, adjust to pH to 8.3; bring volume up to 1L with dH20; store at room temperature. Dilute to 1x with dH20.
- 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.
- 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.
- Destaining 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