A. Protein Precipitation
1. TCA precipitation
Before loading on SDS-PAGE, proteins can be precipitated by mixing 20% TCA with protein samples (1:1, 10% TCA final concentration). Keep samples in ice 20 min. Then centrifuge the samples 10 min at 16.000 x g. The protein pellet is then washed with ethanol/ethyl acetate (1:1) and centrifuged 10 min; 16 000 x g, three times. The final protein pellets is speed-vac dried dissolved in 2X Laemmli buffer at 100 °C for 15 min.
2. Acetone precipitation
Protein samples are mixed with acetone for a final acetone concentration of 80%. Keep samples in ice 20 min. Then centrifuge the samples 10 min at 16.000 x g. The protein pellets is speed-vac dried dissolved in 2X Laemmli buffer at 100 °C for 15 min.
B. SDS Poly-Acrylamide Gel Electrophoresis of Protein
This protocol is part of the following chapter "SDS Polyacrylamide Gel Electrophoresis of Protein" written by John M. Walker, in The Protein Protocols Handbook, Edited by J. M. Walker Humana Press Inc., Totowa, NJ.
1. Preliminary recommendations
To get a high resolution gel:
- you have to minimize the volume of sample to load. For this TCA or acetone precipitation of protein is convenient (see above section).
- The amount of protein loaded should stand in the range of 2 to 50 µg protein. For example, in a case of 1 mm thick gel, I never load more than 20 µl with a maximum of 50 µg. That way, the stacking of proteins is very efficient, this means after migration through the stacking gel, you should obtain a very thin (0.2-0.5 mm) blue band.
- the protein buffer has to be mixed with Laemmli (final 2X)
2. Buffers recipe
|Laemmli 4X (store at RT)|
|Tris-HCl pH 6.8 (250 mM)|
|40% (v/v) Glycerol|
|5% (p/v) SDS|
|0.005% (p/v) Bromophenol Blue|
*Add 10% ß-mercaptoethanol just before use.
Ref: Laemmli, U. K. (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680-685.
The following quantities are adapted for 2 mini-gels of 1-mm thick from the Bio-Rad apparatus.
|Separating gel 12,5%|
|1,875 M Tris-HCl pH 8,8|
|Stock acrylamide (30%)|
|0,6 M Tris-HCl pH 6,8|
|Stock acrylamide (30%)|
|Can be used for 4 runs.|
- Clean with alcohol the glass plates and combs before set-up in the cassette.
- Prepare the stacking gel without the Temed. Degas it 3-5 min, then add the Temed and pour into the cassette until the gel level reaches a position of 0.5-1-cm from the bottom of the comb that will form the loading wells.
- Using a Pasteur pipet, very carefully run UltraPure water down the edge of the plate into the cassette. The water has to spread across the surface of the gel without serious mixing. Wait 10 to 15 min, until a very clear refractive index change can be seen between polymerized gel and overlaying water.
- Prepare the stacking gel without adding the Temed. As previously, degas it 3-5 min. Pour off the water above the separating gel using Whatman paper. Then add the Temed to the stacking gel and poor into the cassette. Quickly, insert the comb in such way you avoid all direct contact between the gel and air, then add with the remaining gel, untill at the edge of the glass (keep in mind that O2 is an inhibitor of the polyacrylamide polymerisation). Put the cassette 30 min at 4 °C, to improve the polymerization of the gel (especially gels wells separating wall). At this stage you can store the cassette for 24-48 hours, wrapped in wet paper and Saran wrap at 4 °C.
- Before using the cassette, carefully remove the comb. With a razor blade, remove all pieces of gel that may obstruct the entry of the gel.
- Load your sample with a loading tip to the bottom of the gel. Fill the tank with the Electrophoresis buffer.
- Connect the power pack to the apparatus, and pass 80 mA through the gel until the sample are totally concentrated in a thin band. Then, increase the power supply to 130-150 mA.
If you want to stain the gel, place it in protein stain reagents for 30-60 min, and wash it several time with the destain reagent. The recipe of the previous reagents are bellow.
Protein stain reagents:
0.1% Coomassie brilliant blue R250 in 50% methanol, 10% glacial acetic acid. Dissolve the dye in the methanol and water component first, and add the acetic acid. Filter the final solution through Whatman N° 1 paper if necessary.
10% methanol, 7% glacial acetic acid
For Western-analysis, go to transfer section.
C. Transfer using semi-dry apparatus
This protocol is part of the following chapter "Protein Blotting by Semidry Methods" written by Patricia Gravel and Olivier Golaz, in The Protein Protocols Handbook, Edited by J. M. Walker Humana Press Inc., Totowa, NJ.
Protein blotting involves the transfer of proteins to an immobilizing membrane.
- I usually use nitrocellulose membrane (Schleicher and Schuell, protran BA 85, pore size 0.4 µm).
- Towbin Buffer: For 1 liter
25 mM Tris 3.03 g
192 mM glycine 14.4 g
10% methanol 100 ml
0.1% SDS 1.0 g
- Adjust volume to 1 liter with dd H2O.
- Note: Do not add acid or base to adjust pH. The buffer will range from pH 8.1 to 8.5, depending on the quality of Tris, glycine, dd H2O, and methanol. Methanol should be analytical reagent grade, as metallic contaminants in low grade methanol will plate on the electrodes.
- Electroblotting apparatus: Trans-Blot SD semidry cell (Bio-Rad). The anode is made of platinum-coated titanium, and the cathode is made of stainless steel.
- Whatman paper, 10 sheets (3MM Chr) by gel.
- The size of the blot and Whatman paper have to fit exactly to the size of the separating gel. This point is important to avoid shortcut of current that would not pass through the gel.
- The membrane, Whatman paper, and gel have to be handle with gloves.
- After the separation of proteins by SDS-PAGE, the stacking gel is cut and discarded, the separating gel is briefly rinsed in distilled water 2-3 min and then equilibrated in Towbin buffer under gentle agitation during 5-10 min. In the mean time, Whatman paper is soaked in Towbin buffer. Wet the anode of the apparatus with Towbin buffer.
- Place the 5 sheets of Whatman paper pre-wetted on the anode (bottom electrode). Then place the nitrocellulose blot on the stack of Whatman paper. Above, place the gel, conveniently oriented (I usually cut the right corner of the gel). At this stage, it's relevant to mark with a pencil the blot at the molecular weight marker positions as the stain of the marker tends to disappear with the process. Place 5 other sheets of Whatman paper pre-wetted. Finally, use a clean tube, and press the stack to roll out all air bubbles. Again, take care that all the elements of the stack fit altogether very well, this is the key of a efficient transfer. See Figure bellow.
- Add some milliliters of Towbin buffer on the top of the stack, wet the cathode with a wet paper towel imbibed with Towbin buffer. Wipe all excess of buffer off the anode, to avoid bridges of buffer that may forms between the 2 electrodes (this is also critical care to have a homogenous and efficient transfer). But, do not wipe too much either. Finally, place carefully the cathode avoiding moving the stack.
- This procedure should not exceed 15 min, because proteins diffuse within the gel and you could lose resolution by being to slow in your handling.
- Connect the blotting unit to a power supply. I usually work at constant power. This means that I correct manually the current that passes through the stack to compensate for the spontaneous decrease of current while the stack tends to dry with time under the established ddp. Usually, I maintain the power at 4-4.5 Watts during 25 min, then I increase it to 5-5,5 Watt during 10 additional min. This gives good transfer for all proteins under 75 k-Da. If your are interested in higher weight protein, increase the 5 Watt step to 20-25 min.
- At the end, cut the power, remove the blot, and dry it on a clean aluminum foil with a hair dryer. This helps to fix the protein on the blot. Do not overdry.
- Proceed to Western-blot treatment (see following section). (You can store the blot at –20 °C after soaking in blocking buffer (TBS, 1% milk) for one hour).
- After transfer the gel can be stained in Coomassie to ensure the completion of transfer.
- The nitrocellulose membrane is blocked in TBS/tween/milk (25 mM Tris-HCl, pH 7.5, 150 mM NaCl, 0,2% (v/v) Tween 20, 1% (w/v) non-fat dried milk) for 1h at RT with gentle rocking.
- Then, incubate the membrane 2h at RT with the primary antibodies (prepared in TBS/tween/milk) and gentle rocking. (This can be done O/N for your convenience)
- Wash 3 times 10 min in TBS/tween/milk (25-50 ml) at RT, gentle rocking.
- Then, incubate the membrane 1h at RT with the secondary antibodies (prepared in TBS/tween/milk) and gentle rocking. (never add the antibodies on the membrane directly without having previously mixed it in the buffer).
- Wash 3 times 10 min in TBS/tween/milk (25-50 ml) at RT, gentle rocking. (The three wash can be do also without milk, in that case the following step is not necessary).
- Rinse 1 time 5 min in TBS/tween WITHOUT milk at RT, gentle rocking (~50 ml).
- For alkaline phosphatase staining, incubate the membrane 5 min at RT and gentle rocking in 25 ml of revelation buffer (100 mM Tris-HCl, pH 9.5, 100 mM NaCl, 5 mM MgCl2). Then add the substrates: 100 µl BCIP (25 mg/ml dissolved in 100% dimethyl formamide); 100 µl NBT (25 mg/ml dissolved in dimethyl formamide/H2O : 70/30). 15 min of incubation is an average time that you may modify accordingly to your result.
- Wash thoroughly with water to stop the reaction. Dry the blot with a hair dryer (do not over-dry). Store the membrane in the dark for example between Whatman sheets in your lab-book (formazan staining is light sensitive).