Running an SDS-PAGE Gel:

Note that the following uses pre-cast gels and pre-made running buffer, see accessory protocols [NotDoneYetDudez] for casting gels and making your own running & loading buffers

Supplies:

  • ~1L of 1x Invitrogen MES PAGE Running Buffer (supplied at 20x, Life Tech. Cat. No. XXXX)
  • 1 or 2 precast acrylamide gels (I like the 4-20% gradient gels, e.g. Life Tech Cat. No. XXXX)
  • 4x SDS-PAGE Running Buffer (Life Tech Cat. No.)
  • Gel Running Chamber (Life Tech Cat. No. XXXX)
  • DC Power Supply
  • Prestained Protein Ladder (e.g. Pierce PageRuler Plus, Pierce Cat. No. XXXX)
  • SimplyBlue SafeStain Coomassie staining solution (Life Tech Cat. No. XXXX)
  • Thermocycler or 95degC Heat Block

Preparing the Samples:

  • For protein solutions (e.g. column flow-through, cleared lysate, purified protein):
    1. Add 1/3 vol. 4x SDS-PAGE Sample Loading Buffer and pipette up and down to mix
    2. Incubate sample at 95degC 5min
      • I like to do this step in the thermocycler; prepare < 50uL of sample + loading buffer in a 0.2mL PCR tube or strip and incubate at least 5min @ 95C

  • For E. Coli cultures (e.g. to check protein expression):
    1. Remove a small amount of culture (25uL is good) DON'T spin down
    2. Add 1 vol. of 4x sample buffer directly to the culture
    3. Incubate 5min @ 95degC in the thermocycler
    4. Dense cultures can become viscous and nearly impossible to load after boiling; if this is the case, repeat above steps with 1-3 more volumes of sample buffer, or dilute culture 2-4x with water

  • spin tubes briefly to remove condensation from cap
  • Note that you don't need to boil the pre-stained ladder

Setting up the Gel Rig, Loading, and Running:

  1. Prepare 1x running buffer (50mL of 20x in 1L ddH2O is usually enough for mini gels)
  2. Open gel packages
    • MAKE SURE TO REMOVE TAPE STRIP FROM BOTTOM OF GEL OR IT WON'T RUN
  3. If running 2 gels, put one on either side of the white electrode assembly, with the short plate facing inwards (gels should be 'facing' one another)
    • If only running one gel, use the plastic buffer dam in place of the 2nd gel (writing faces inwards)
  4. Remove plastic combs from the tops of the gels
  5. Pour running buffer into top chamber (between gels) until buffer level is higher than the top of the (shorter) inner gel plate (i.e. until buffer covers the wells). Leave for a minute or two to make sure there are no leaks (buffer level doesn't drop).
  6. Fill the outer chamber until ~3/4 of the gel is covered.
  7. [Optional, but recommended] Use a 1mL pipet to wash some running buffer into wells to clear out any gel fragments or unpolymerized acrylamide
  8. Load samples into wells with a P20 tip or gel-loading tip
    • 5uL is usually sufficient for a 1mm gel; I wouldn't load more than 10uL unless you know you have a very dilute sample
    • Load 2-5uL of Prestained Ladder
  9. Attach lid and run gel!
    • 1hr at 150V is usually sufficient
    • If running for a western blot, run until dye front has passed completely out of the gel

Staining (with SimplyBlue Coomassie):

  1. Remove gels & break open plastic cassette by using a metal scoopula to pry apart the plates along the edges
  2. carefully remove top (short) plate
    • lift from bottom of gel to prevent gel sticking to plate
  3. use a razor to cut along edges of the gel and remove the comb
    • cut by pushing directly down into the gel; if you run the razor horizontally along the edge the gel will tear
    • If staining multiple gels, cut off a corner to tell them apart
  4. CAREFULLY lift the gel out of the cassette (very easy to tear, esp. 1mm gels!). Transfer to a tupperware container or pipet tip box lid
    • Can stain 2 gels in one container
  5. Repeat 3x:
    • Add 100mL of ddH2O, and microwave one minute at full power
    • Gently shake on an orbital shaker (~60RPM) for 1min
    • Pour off water into sink
      • hold onto corner of gel, easy to lose it down the drain
  6. Add 25mL of SimplyBlue Coomassie stain to the gel(s) and microwave until stain just starts to boil (usually < 30s)
    • use 35mL for thick gels or to do two thin gels at once
  7. Cover and gently shake (<60rpm) at RT for 5 minutes
  8. Pour off stain (can dump down drain) and add 100mL diH2O
  9. Shake 10min at RT
  10. [OPTIONAL (though I almost always do it)]:
    • Pour off H2O and add same volume of 20% NaCl, then shake 5min RT
    • Gel will be stable in this solution at 4C for a few days at least
  11. Image gel on the WEL gel doc using the COOMASSIE BLUE protocol with the white-light screen in place.
    • Gels can be disposed of in regular trash


This topic: Lab > WebHome > ProtocolList > ProtocolsRunningSDSPAGEProteinGels
Topic revision: r1 - 28 Jun 2016 - 15:46:11 - Main.ColinBrown