SDS-PAGE, Transfer, and Western Protocols

SDS-PAGE

  1. Clamp a glass plate to the empty side of gel apparatus if running only one gel. Put gel seal around gasket on side you are using, choose a pre-poured gel and clamp it into position, white plate facing inward.

  2. Prepare stacking gel, and pour over gel with 10 ml pipet. Need about 3.3 ml per gel.

  3. Insert comb between plates. Avoid bubbles.

  4. Let set ~20 minutes to harden.

  5. Fill both chambers with 1X SDS Buffer. Remove comb. If necessary, blow out wells with pipet tip.

  6. Load 20-30 ul samples using gel loading pipet tips. Amount of sample depends on the size wells you chose.

  7. Run at 95-105 Volts for ~2 hours, or 35 mAmps per gel for ~1 hours. Check that your samples do not run off the gel. They will if you leave it on too long and do not check it.

  8. When done, turn off power, unplug and discard buffer.

  9. (optional) If the proteins are not to be transfered to nitrocellulose, then the gel can be stained. For Coomassie staining, place the gel in Coomassie stain for 2-24 hours. Rinse gel with water and place in destain until background is clear and bands are visible.

TRANSFER

  1. Dilute 40X WTB buffer to 1X WTB and 10% methanol in distilled water. For 1 transfer apparatus you need about 1 to 1.5 liters. Buffer can be recycled one or two times. 25ml 40X WTB in ~800 ml distilled water. Add 67 ml methanol. Bring up to 1 Liter with distilled water. (Dilute WTB in distilled water before adding methanol or else the methanol will begin to precipitate the 40X WTB).

  2. Place sandwich apparatus with the sponges in a pan, white side down. Cover with buffer, let sponges soak up the buffer and remove bubbles. Put down 3 pieces of whatman paper cut slightly larger than the gel. Put down 2 pieces nitrocellulose paper cut slightly larger than the gel. Put gel on top. Put another 3pieces of whatman paper on top. Place a stir bar in the gel box. Fill box with buffer. Close the sandwich, squeeze bubbles out, and place in box. Put leads on black wire on black side of sandwich, red wire on white side of sandwich. (Protein and DNA run to red (+) terminal). Put on stir plate and plug in to power supply in cold room. Run for 2 hours (use the timer if available) at 150mA current, Voltage set as high as it can go. If you use the timer, it will shut-off by itself and can be left overnight.

  3. Remove nitrocellulose from box. Place TOP piece (piece closest to the gel) in TBST with 5% evaporated milk. Place 2nd piece of nitrocellulose in container with TBST and ~5 drops india ink. Incubate on shaker at least O/N. Dry and you will see your bands. If destaining of nitrocellulose is necessary, destain in TBST.

WESTERN

  1. Place nitrocellulose in TBST with 2% milk and 1-10ul of primary antibody. Incubate at room temperature for 2 hours.

  2. Rinse nitrocellulose 3 times with distilled water. Wash nitrocellulose twice in TBST at 5 minutes per wash.

  3. Place nitrocellulose in TBST with 2% milk and 1-10ul of secondary antibody. Incubate at room temperature for 2 hours.

  4. Rinse nitrocellulose 3 times with distilled water. Wash nitrocellulose twice in TBST at 5 minutes per wash.

  5. Incubate nitrocellulose in ECL reagents for 1-2 minutes.

  6. Place nitrocellulose on plastic wrap and blot dry with paper towel.

  7. Expose nitrocellulose to film.


Recipe for 16% SDS-PAGE gels

TO MAKE 10 gels (80 ml):

  • 17.4 ml distilled water
  • 42.6 ml 30% Acrylamide
  • 20 ml lower buffer
  • 320 ul 10% ammonium persulfate (APS)
  • 48 ul TEMED

Recipe for 8% SDS-PAGE gels

TO MAKE 10 gels (80 ml):

  • 38.7 ml distilled water
  • 21.3 ml 30% Acrylamide
  • 20 ml lower buffer
  • 320 ul 10% ammonium persulfate (APS)
  • 48 ul TEMED

Recipe for Stacking gels

TO MAKE 10 ml:

  • 6.2 ml distilled water
  • 1.3 ml 30% Acrylamide
  • 2.5 ml upper buffer
  • 100 ul 10% ammonium persulfate (APS)
  • 20 ul TEMED

LOWER BUFFER
(pH 8.8, 1.5 M Tris, 0.4% SDS)

TO MAKE 500 ml:

  • 91.0 g Tris
  • 20 ml 10% SDS stock solution

Add Tris to distilled H2O to equal about 400 ml. Adjust pH to 8.8 with HCl. Then add SDS. Bring up to 500 ml with distilled water.

UPPER BUFFER
(pH 6.8, 0.5 M Tris, 0.4% SDS)

TO MAKE 500 ml:

  • 30.5 g Tris
  • 20 ml 10% SDS stock solution
  • 1-3 ml 0.01% Bromophenol Blue stock solution

Add Tris to distilled H2O to equal about 400 ml. Adjust pH to 6.8 with HCl. Then add SDS. Bring up to 500 ml with distilled water.

ELECTROPHORESIS RUNNING BUFFER
(250 mM Tris, 1.92 M glycine, 1% SDS))

TO MAKE 1 L (10X):

  • 30.3 g Tris
  • 144 g Glycine
  • 10 g SDS

Dissolve in distilled H2O to a final volume of 1 Liter. Requires Heat. Wear a mask handling SDS.

COOMASSIE STAIN

TO MAKE 500 ml:

  • 0.5 g Coomassie Blue
  • 10 ml distilled water
  • 30 g NH4SO4
  • 500 ml 2% H3PO4

In 50 ml conical tube dissolve 0.5 g of purified, dried coomassie blue in 10 ml distilled distilled H2O. Nutate at room temperature from 2-24 hours.

In a beaker, dissolve 30 g NH4SO4 in 500 ml 2% H3PO4. Dissolve completely.

Add the coomassie blue mixture to the beaker and mix. Store at room temperature in a dark bottle (light sensitive).

WESTERN TRANSFER BUFFER (40X)
(264 mM Tris, 2 M Glycine, 0.4% SDS)

TO MAKE 500 ml:

  • 16 g Tris
  • 75 g Glycine
  • 2 g SDS

Wear a mask while handling the SDS. Requires Heat to get SDS into solution.

TBST (20X)
(200 mM Tris, pH 8.0, 3 M NaCl, 1% (v/v) Tween 20)

TO MAKE 1 L:

  • 24.4 g Tris
  • 175.2 g NaCl
  • 10 ml Tween-20
  • 4 g NaN3 (optional)

Adjust pH to 8 with HCl BEFORE adding detergent. Usually takes about 12 ml HCl for 2 Liter recipe.