SDS Gel Electrophoresis and Western Blot Protocol

Biochemistry, Molecular Biology, and Cell Biology Protocols >> SDS Gel Electrophoresis and Western blotting Protocol

RECIPES:

when working with proteins, try to use nanopure water

10% SDS
10g SDS
up to 100mL water

Buffers for Making Gels
Lower Buffer (for the separating gel)
1.5M Tris 36.4g pH to 8.8 with 6M HCl until nearing desired pH
0.4% SDS 8mL of 10% SDS
up to 200mL with water

Stacking (Upper) buffer (for the stacking gel)
requires fair bit of HCl so start with ~70mL water when add Tris
0.5M Tris 6.06g pH to 6.8 with 6M HCl until nearing desired pH
0.4% SDS 0.4g
up to 100mL with water

30% Acrylamide*
Acrylamide 30g
Bis 0.8g
up to 100mL with water
wrap aluminium foil around bottle because it is light sensitive

Making SDS Polyacrylamide Gels*
(once the last ingredient, ammonium persulfate, is added, the gel will begin to polymerise)
Separating (lower) Gel - 10% acrylamide (change the proportions of water and acrylamide if different from 10% acrylamide)
Lower buffer                  1.9mL
Water                              3.1mL
Acrylamide                     2.5mL
TEMED                           10uL (add TEMED in the fume hood)
Ammonium Persulfate 20uL
Total                                 7.5mL

*Note that acrylamide in its unpolymerized form is a potent neurotoxin and that gloves must be worn for making gels, setting up the tank for running the gel, and during transferring of the gel

Stacking Gel (for all percentages of lower gel, use this upper gel)
Upper Buffer                     1.25 mL
Water                                  3.25 mL
Acrylamide                         0.5 mL
TEMED                               10uL (add TEMED in the fume hood)
Ammonium Persulfate    20uL
Total                                     5 mL

10x Electrophoresis Running Buffer (for the tank)
Tris 121.4g
Glycine 567g
SDS 40g
Water up to 4L

pH 8.3 without adjustment

4x SDS Loading buffer
Glycerol 4mL (put an empty flask onto balance and weight out glycerol by pipetting)
SDS 0.4g
Upper buffer 5mL
up to 10mL with water
· heat while stirring to dissolve the SDS (don’t make the solution boil when heating), and add enough bromophenol blue to make the 1x solution dark enough for easy monitoring when running on a gel
· store at room temperature

1x Coomasie Blue Dye (for staining gels that are not used for transferring, or for ensuring that no protein is left after transfer)
Isopropylalcohol 25% 500mL
Acetic Acid 10% 200mL
R250 Coomasie Blue 0.025% 0.5g
Water 1.3L
(destain is 10% Acetic Acid)
· stain 4 hours to ensure all the protein is stained
· destain long enough so that the background is clear (a piece of foam or paper towel may be added to the solution to speed up the destaining)

10x Towbins Transfer Buffer (for semi-dry transfer of protein from gel to blot)
250mM Tris 15.1g
1.92M Glycine 72.0g
Water up to 500mL
pH 8.3 without adjustment

10x Phosphate Buffered Saline—PBS (for removing methanol from blots)
NaCl          1.37 M for 10x
KCl 2g       27mM for 10x
Na₂HPO₄80mM for 10x
KH₂PO₄   20mM for 10x
water to 1L
pH of the 1x PBS should be 7.4

PBS Tween—PBST (for washing blots)
PBS 2L
Tween-20 (Brown bottle) +1mL by pipetting it in with pipettor or plastic bulb
stir half an hour

Amido Black Stain (for viewing all proteins non-specifically after blotting)
Methanol 40mL
Acetic Acid 10mL
Amido Black 10B 0.1g
Up to 100mL

Western Blot protocol

Making the gel
1. Use alcohol and Kimwipes to wipe the glass, and then set up the rest of the apparatus
2. For small proteins, use a higher percentage gel
2. Add all ingredients except ammonium persulfate; before adding ammonium persulfate, if the weather is cold in the room, you may wash the solution with hot/warm tap water to help the polymerization
3. Invert tube gently after adding all components
4. Add it up to the correct mark
5. Add t-butanol
6. Remove t-butanol, wash with water, and wipe with filter paper; put in comb
7. Make upper gel and gently shake 2x
8. Add upper gel
9. Remember to keep tubes to check if they’ve polymerised (cap the tube after pouring and see if the material in the tube has polymerised)
10. Label the lanes of the upper gel with a marker if required

Preparing and Loading Samples
1. Set up the gel in the tank and make sure the buffer isn’t leaking (if it is, it means that the glass plates are not slightly protruding up from the screws on both ends)
2. Pour in some running buffer to prevent drying of the gel
3. For each sample, use 1/3 the volume of 4x loading dye (e.g. if have 30uL of sample, add 10uL of 4x loading dye)
4. If running samples in reduced form, add 5% beta-mercaptoethanol in the fumehood
5. once the loading dye has been added, do no place on ice, or the SDS will precipitate out

Running and transferring the gel
1. Check no leaking and it’s running
2. Set for anywhere from 20-30mA, and all the voltage it needs
3. Keep an eye on it to ensure there is current
4. Cut out filter paper and nylon—eight sheets of filter paper for each blot
5. Mark the front of the nylon (make a cut)
FOR NITROCELLULOSE BLOTS DON’T WET WITH METHANOL, OR ELSE IT’LL DISTINTEGRATE IN YOUR HANDS
6. Wet the nylon with methanol
7. Immerse gel, filter paper, and nitrocellulose in transfer solution
(20% methanol + Towbins 1x); use a lower percentage of methanol for larger proteins (e.g. 7%)
8. Proteins in SDS are negatively-charged, so place sandwich as follows
(flatten with each layer except for gel layer because gel is fragile)
3 pieces filter paper
nylon
gel (proteins in SDS are negative)
3 pieces filter paper
9. For transfer conditions, check with manufacturer. We normally set at 20V, 0.300A, 20 minutes (time varies—larger proteins require longer time)
10. Rinse blot with distilled water and hang to dry
11. wipe the transfer apparatus with plenty of water
12 to dry the transfer apparatus more quickly, wipe with some towels, and then fan the remaining liquid with a piece of paper towel

Blotting
1. Wet a dried blot with methanol if the blot is made with PVDF, or soak in PBS if blot is made from nitrocellulose (note that methanol is toxic, so wear gloves and don’t inhale it)
2. Rinse ~4x with PBS to get rid of the methanol
3. Place the blot face up for blotting
4. 1% skim milk (made in PBS) 10 min. (some proteins require a higher percentage of skim milk e.g. 5% and incubation in the cold room for longer e.g. 30min.)
5. 10mL total--Primary antibody diluted to the appropriate concentration in PBS (+0.2% milk) 1hr
6. 3x 10min. each PBST
7. 1:5000 dilution of secondary (choose the right secondary—either goat anti-rabbit, or goat-anti mouse), 30 minutes
8. Wash ~5x PBST 5-10 minutes each

Viewing
1. mix equal amounts of solution A & B of ECL (~1.5mL each of A & B)
2. rock manually for 1 min.
3. wrap Saran Wrap around blots
4. take off gloves when handling film
5. turn off all lights (can leave a dim red light on) when removing film from the box, and place remaining film back into box before dealing with the newly-removed piece of film
6. place film on top of blot in the film cassette, making a note of the orientation of the film
7. expose for 15sec -2 minutes, and if required, use another film and expose for anther 20 min.
8. develop

Nonspecifically staining the blot
1. rinse blot with distilled water several times to remove the ECL
2. invert amido black bottle a few times, and pour some into a petri dish with the blot in it
3. rock blot a few times
4. pour amido black back into bottle
5. rinse with distilled water several times to destain the blot
6. hang blot to dry and place in notebook
If blot turns dark blue irreversibly, dry or put in PBST and then 100% methanol