Difference between revisions of "Lidstrom: SDS-PAGE"

From OpenWetWare
m (Running Buffer:)
m (Running/Electrode Buffer:)
Line 140: Line 140:
* 10x SDS-PAGE (1 L) (BioRad catalog #161-0732) = 250 mM Tris, 1.92 M glycine, 1% SDS, pH 8.3  
* 10x SDS-PAGE (1 L) (BioRad catalog #161-0732) = 250 mM Tris, 1.92 M glycine, 1% SDS, pH 8.3  
***Tris base  30.30 g '''DO NOT USE Tris-HCl'''
***Tris base  30.30 g     '''DO NOT USE Tris-HCl'''
***Glycine  144.10 g  
***Glycine  144.10 g  
***SDS  10.00 g  
***SDS  10.00 g  

Revision as of 18:47, 14 November 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
  • 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.
  • Use pipette to put gel mix into the gap between the plates
  • Insert the comb being 'careful not to trap any bubbles'
    • It is much easier to avoid bubbles if you fill the space with enough solution that it spills over the top as you put the comb in.
  • 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.

Consolidated/Pictorial Protocol

SDS casting protocol

Sample Prep

  • use 5-8 or even up to 20 ug protein per well. (for Mini Protean with 10 well comb)
  • If biomass is not limiting, prep a significant excess of protein, which will allow you to re-run if your gel turns out poorly for any reason.
  • If protein normalizing:
    • lyse cultures by sonication. Use 20 1 second pulses while tube is in ice, let the samples rest 5 minutes on ice, then sonicate again. Wipe sonicator stick between uses. Don't touch the tip while it is on.
      • sonication allows you access to un-dyed lysed sample, unlike boiling with sample loading buffer as is described below.
    • do Pierce BCA BSA assay to determine concentration. Run each sample with dilutions: 1, 1/10, 1/20, etc.
    • Boil samples with sample buffer + beta-mercaptoethanol
    • Optional: check efficacy of lysis with light microscopy (people usually don't do this.)
    • Centrifuge all extracts extensively (20,000 x g for 15 min at 15°C) to remove any insoluble mater
      • You can run gels with the soluble, insoluble, and/or crude (not centrifuged) lysates.
  • If not normalizing:
    • Boil 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)
    • consider centrifugation to remove insoluble??
    • heat sample to 37oC to solubilize precipitated SDS??
  • Load 12-15 uL, absolute max is 30 uL for 10-well comb in mini-protean
    • less for the skinny wells
  • If gel is overloaded or underloaded, run a new gel with a different amount of dyed & boiled lysate

Consolidated/Pictorial Protocol

SDS-PAGE sample prep and loading

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 you will be able to determine the orientation of your gel after it is stained. Asymmetry is good!
  • 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 ~15 min, then 200 V for ~ 20+ min.
    • Note: ladder looks blurry while running through the stacking gel; don't be alarmed unless it still looks blurry in the resolving gel.
    • 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 the protein doesn't run 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

  • 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
  • taking the gel off the casting stand before it has polymerized
    • entire sample will leak out
  • 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.
  • not having the lid to the running unit on all the way. --> poor electrical contact & blurry bands
  • running the gel without enough buffer between the gel plates
    • will cause the gel to run terribly (unusable)
    • avoid by making sure the plates are well-seated


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.
  • Can add up to 8M urea for really hydrophobic proteins

Running/Electrode Buffer:

  • 10x SDS-PAGE (1 L) (BioRad catalog #161-0732) = 250 mM Tris, 1.92 M glycine, 1% SDS, pH 8.3
    • Mix:
      • Tris base 30.30 g DO NOT USE Tris-HCl
      • Glycine 144.10 g
      • SDS 10.00 g
      • diH2O to 1 L
    • Do not adjust the pH (~pH 8.3)
  • 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.
  • It is best practice to not re-use electrode buffer. Our lab does, however, re-use the buffer ~ 4 (but sometimes up to 10) times.

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


PageRuler protein gel legend

Other ladders:

BioRad protein standards

Combs/Loading Volumes

  • We currently only have the mini-protean gel running boxes. -JM 10/2012
SDS combs and loading volumes


What do all of the reagents do?

  • SDS
    • In loading buffer and often in gels (not necessary to include in gels; can be used in sample buffer)
    • polymerize acrylamide
  • glycine
    • carry charge in the opposite direction as the negatively-charged SDS-covered proteins
  • methanol
    • in wash buffer
  • glycerol in loading buffer: helps sample sink
  • Bromphenol Blue

Is it important to degas my water + buffer + acrylamide mix before adding the APS and TEMED, as the manuals recommend?

  • "We recommend that you degas the solutions to get rid of nitrogen. Sometimes, the nitrogen may come out of solution and form bubbles in your gels. " -BioRad customer support 10/26/2012 JM
  • "Proper degassing and filtering of the casting solution is critical for both reproducibility of the polymerization (oxygen removal)" (link)
    • This manual suggests the consequence is poor polymerization. If you don't experience poor polymerization, maybe you don't need to worry about degassing...?
    • Also note this is not the justification customer support provided. (see above)

Why shouldn't we overlay the gels with butanol or isoporpanol as they polymerize?

  • "We do not recommend using butanol/isopropanol because these may degrade the glue on the spacer plates and the plastic of the casting frame." -BioRad customer support 10/26/2012 JM

Should I soak my gel or run them empty before use?

  • They say no: "I have not heard that soaking the gels in water will improve the run. Talking with my colleagues, we thought it would have been detrimental because the tris and chloride ions will diffuse out during this time and in theory, should have made your gel run poorly."
  • JM had a gel that was only loaded with two lanes of ladder run funny. The ladder bands ran fine until 1/2 way through the gel, after which they halted and stacked on top of one another. Immediately after, I could see a 1 cm band in the gel that refracted light differently. This band disappeared after a while of soaking, leading me to believe it was a buildup of one of the buffer compounds. I soaked this gel overnight in water, loaded ladder in an unused adjacent well, and it ran perfectly without bunching up. The ladder ran fine until it reached ~ 1/2 way down, where it bunched up. A ~ 1 cm horizontal strip of the gel refracted light differently, but this band went away over time. It was repeatable with gels in the same batch, and with a batch Amanda prepped (though hers were less dramatic).
  • We currently believe it is best to soak gels overnight or run them in buffer for ~ 1 hour at 200V before use. BioRad tech support suggested that if we chose to soak (which they don't officially recommend), we should do so with You can use 0.375 M Tris, pH 8.8. This is the concentration of the storage buffer used in BioRad's precast gels. -JM 10/29/2012
  • Potential problems with soaking or pre-running gels:
    • You might ruin the stacking nature of the gel by alterning the buffer within the acrylamide matrix. This essentially converts your gel to a continuous buffer gel system, known to give more blurry bands.

Is it ok to re-use my electrode buffer?

  • Many people re-use it up to ~ 20 times, however, you should know that the manuals recommend single use and understand why. This may help.

How long can I store my acrylamide gels?

  • "Tris-HCl resolving gels are prepared at pH 8.6–8.8. At this basic pH, polyacrylamide slowly hydrolyzes to polyacrylic acid, which can compromise separation. For this reason, Tris-HCl gels have a relatively short shelf life. In addition, the gel pH can rise to pH 9.5 during a run, causing proteins to undergo deamination and alkylation. This may diminish resolution and complicate postelectrophoresis analysis." (reference)
    • note: commerical Tris gels have a shelf life of 6 months to 1 year. Amanda & Ceci don't keep hand-poured gels for more than a few weeks. (JM 11/2012)
    • It is possible the commercial gels have other means of extending shelf life.

The two types of Coomassie Blue dyes

  • useful link
  • 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.

Other Tips

Excess salt in SDS-PAGE samples causes fuzzy bands and narrowing of gel lanes toward the bottom of the gel

  • If the ionic strength is very high, no bands will appear in the lower part of the gel (a vertical streak will appear instead) and the dye front will be wavy instead of straight. Deionize any sample with a total ionic strength over 50 mM using columns such as Micro Bio-Spin™ columns, which contain 10 mM Tris at a pH suitable for SDS-PAGE. (source)

Success or failure of any protein analysis depends on sample purity.


  • Interfering substances that can negatively impact SDS-PAGE include salts, detergents, denaturants, or organic solvents (Evans et al. 2009). Highly viscous samples indicate high DNA and/or carbohydrate content, which may also interfere with PAGE separations. In addition, solutions at extreme pH values (for example, fractions from ion exchange chromatography) diminish the separation power of most electrophoresis techniques. Use one of the following methods as needed to remove these contaminants:
    • Protein precipitation — the most versatile method to selectively separate proteins from other contaminants consists of protein precipitation by trichloroacetic acid (TCA)/acetone followed by resolubilization in electrophoresis sample buffer. A variety of commercial kits can simplify and standardize laboratory procedures for protein isolation from biological samples
    • Buffer exchange — size exclusion chromatography is another effective method for removing salts, detergents, and other contaminants

Polyacrylamide is oxidative, and so disulfide bonds may be re-formed after proteins enter the gel

  • also, oxidation of agents used to reduce any disulfide bonds in the original sample may introduce artifactual new components in the gel pattern. (source)

SDS can form micelles

  • The SDS concentration is greater than the critical micelle concentration (cmc), so SDS is present as micelles. Micellar SDS bound to the tracking dye unstacks at high gel concentrations in disc electrophoresis; so a tracking dye that marks the moving boundary front at low gel concentrations fails to do so at high concentrations (source)

KCl causes SDS to precipitate

  • If you samples contain KCl you should dilute them or methanol precipitate them and resuspend them in 1X sample buffer. With low concentrations of KCl (<200 mM) you can run them on the gel but you should loaed every lane with sample buffer containing the same concentration of KCl (even if they are blanks). This will help the gel run a little less anomalously. (source)