SDS-PAGE Gel Staining

Blue Silver : most sensitive method

10% phosphoric acid

10% ammonium sulfate

0.12% G-250

20% methanol or EtOH

How to make:

make a solution containing 10% phosphoric acid,  10% ammonium sulfate, 0.12% G-250. Then while stir slowly, slowly (drop by drop) add methanol

To 100 ml water add phosphoric acid (enough to obtain 10% in the final 1L).

• Add 100 g ammonium sulphate.
• Add 1.2g Coomassie Blue G-250.
• Add 116 mL of 85% o-phosphoric acid
• Add water to 800 mL.

Dissolve by stiring for 1 hour. then:

• Very slowly using peristaltic pump or flow rate adjusted column: add 200 mL of 100% methanol.

The critical point lays in the mode of addition of methanol, which facilitates the staining. Methanol addition to the solution of 10% phosphoric acid, and 10% ammonium sulfate is a critical step. It should be added extremely slowly (ideally in a 'drop-by-drop' manner, i.e., in small amounts and not in one step), under continuous and vigorous stirring in order to preserve the micellar colloidal suspension of the dye. This improves sensitivity. Indeed, the final physicochemical conditions of the solution are not the same in either case. The physical state of the solution can be easily inferred by the color of the solution: it will be a dusty greenish-blue when addition has been performed slowly but light blue if addition has been carried out incorrectly (too quickly). In the latter case, the homogenous blue color denotes dispersion of the micelles, with loss of the improved sensitivity. When addition of methanol is done in a one-step procedure instead of adding it 'drop by drop' as stated above with extremely gentle stirring (being perhaps the only time-consuming step, to be done the day before need) colloidal micelles are suddenly dispersed and the color of the solution shifts from a greenish dusty deep blue to a brilliant homogenous blue (typical of the formation of unprotonated anionic species, and much less efficient in complexation with proteins). When micelles are dispersed, the sensitivity of the method drops from 1–5 ng down to 10 ng or worse. The state of the micelles is pivotal, therefore any care must be taken in avoiding micelle solubilization

Sensitive staining:

1. Fix gel in 50% ethanol and 2% phosphoric acid.
2. Wash the gel for 2x 20 minutes in ddH2O.
3. Add staining solution and stain overnight or longer.
4. Rinse the gel with ddH2O and destain with ddH2O using microwave.


Destaining: water

rinsing and fixation in 20% methanol

Protocol 2

Blue Silver staining method uses Coomassie Brilliant Blue G250 for the staining reaches sensitivity upto 1ng. The method was originally developed by Neuhoff and further modified by Righetti et al, 2004.

Procedure
1) Following the electrophoresis fix the resolved protein with Acetic acid: Methanol: Water 1:3:6. Alternatively One can use 10% Phosphoric acid.
2) To the CBB G250 solution add 20% Methanol just prior to use with constant stirring to allow the formation of colloid effectively.
3) Add the above prepared solution to the gel and keep on a gel rocker with constant shaking.
4) Protein bands take up bright blue stain while the background remains faintly stained with this procedure.
5) Briefly rinse the gel with Tris-Phosphate solution pH6.8.
6) Stabilize the protein-CBB G250 complex with 20% ammonium sulphate solution for 30min.
7) Rinse with water and store the gel indefinitely in distilled water.

Recipes
CBB G250 solution - 0.12% CBB G250 in 10% Phosphoric acid, 10% ammonium suphate and 20% methanol.
The CBB G250 solution is prepared as follows:
to a water solution (1/10 of the final volume) the desired amount of phosphoric acid is added, so that, in the final volume, its concentration will be 10%; to this, add the required amount of ammonium sulfate (in powder), calculated to obtain a final concentration of 10%. When the ammonium sulfate has dissolved, add enough Coomassie Blue G-250 (in powder) to obtain a final concentration of 0.12%. When all
solids have dissolved, add water to 80% of the final volume. To this solution, under stirring, add methanol to reach a 20% final concentration. This stock dye solution should be kept in a brown bottle and is stable at room temperature for almost 6 months.

Colloidal Coomassie

Colloidal Coomassie staining with CBB G-250
1. Staining solutions
Note: for the preparation of the staining solutions, use chemicals with high quality such as analytical grade (p.a.) and water of high purity as you get from Millipore systems (Milli-Q water).

Coomassie solution:		ad 2000 ml H2O
0.02 % (w/v)	CBB G-250	0,4 g
5 % (w/v)	aluminum sulfate-(14-18)-hydrate or ammonium sulfate	100 g
10 % (v/v)	ethanol (96%)	200 ml
2 % (v/v)	orthophosphoric acid (85 %)	47 ml

Note: for the preparation of the staining solution, the sequential addition of the components in the following order has to be maintained:

1. first dissolve aluminum sulfate in Milli-Q water
2. thereafter add ethanol, homogenize, and mix CBB G-250 to the solution
3. as recently as the solution is completely dissolved, add phosphoric acid (the incorporation of the acid to the alcoholic media lets the Coomassie molecules aggregate into their colloidal state)
4. finally fill up with Milli-Q water

Note: the final staining solution has a dark green-bluish appearance and is full of particles swimming around:

• Don't filtrate this solution!
• If you changed the order of preparation, you would get a more violet-bluish solution that is less sensitive.

Destaining solution:		ad 2000 ml H2O
10 % (v/v)	ethanol (96 %)	200 ml
2 % (v/v)	orthophosphoric acid (85 %)	47 ml

2. Staining procedure

• After second dimension separation, carefully remove the gels from the glass plates and transfer them into a staining dish filled with Milli-Q water.
• Wash the gels three times with Milli-Q water for 10 minutes on a horizontal shaker (insufficient washing causes poor sensitivity because the remaining SDS on the gels disturbs the bounding of the dye to the protein).
• Shake the Coomassie solution before use to disperse the colloidal particles evenly.
• Incubate the gels well covered with the Coomassie solution by agitation on a shaker for 2-12 hours. This staining generates marginal background so you can observe the staining progress in between.
  Note: after 10 minutes you can see first protein spots appearing, within 2 hours of incubation about 80% staining to its maximum level is completed. For nearly 100% staining, overnight incubation is recommended. In some cases, when the amount of protein to be stained is huge and the solution turns into a bright blue, a refresh of the staining solution is necessary.
• After the staining procedure remove the Coomassie solution and rinse the gels twice with Milli-Q water.
  Note: you can reuse the staining solution as long as particles are still remaining, otherwise discard it (keep in mind that your laboratory may have special regulations for waste disposal).
• Remove sticking dye particles from the staining dish with a lint-free paper towel and destain for 10- 60 minutes.
  Note: You can also wash the gels just with Milli-Q water but this we recommend only for preparative gels because there will last an irregular, weak Coomassie film on the gels that will arise during the scanning procedure.
• Finally rinse the gels twice with Milli-Q water: the gels will reach their original thickness (the alcohol-acid media makes the gels shrinking) and neutralization in water additionally enhances Coomassie's color intensity.

Colloidal CBB

Colloidal CBB staining came later on, thanks to the attempts to reduce the background staining. Diezel et al. observed that CBB-G in 12.5% Trichloroacetic acid is converted into a colloidal[23] hindering dye entry into the gels, therefore giving a low background. Neuhoff et al. was the first to develop a staining method using colloidal CBB-G in the presence of ammonium sulfate and phosphoric acid, demonstrating the benefits of adding alcohol to the staining solution.[24] This is allowed the detection of weak spots.[19,20] The addition of methanol in the classical staining solution (20% methanol, 1.6% phosphoric acid, 8% ammonium sulfate, 0.08% CBB G-250), intended to shift the equilibrium from the colloidal form to dispersed dye, and facilitated the staining;[24] however, this introduced a main critical point, as will be discussed below. The principal characteristics that make colloidal CBB an excellent staining method are its ease of use, extremely low signal-to-noise ratio and its sensitivity.[16] Colloidal particles are in equilibrium with dispersed dye in solution. Protein staining (and low background) is achieved when the free dye enters the gel, while the colloidal dye particles are excluded. Lately, Candiano et al.,[25] Wang et al. [26] and Pink et al. [27] proposed different protocols aiming to overcome the limit of detection of CBB staining, still requiring approximately 10 ng of protein in a band. 'Blue silver' was developed by Candiano et al. as a modification of Colloidal CBB increasing dye by 20% and phosphoric acid by fivefold (staining solution: 20% methanol; 10% phosphoric acid; 10% ammonium sulfate; 0.12% G-250).[25] The lower pH, causing protonation of Asp and Glu residues, is thought to promote more efficient hydrophobic binding to aromatic and hydrophobic residues, allowing a more intensive staining. Blue silver is more sensitive than the other colloidal CBB-G stains, with a detection limit approaching 1 ng for BSA. The possibility of destaining 'blue silver' exclusively with ultrapure water is also advantageous as it is low in cost and minimally affects stained bands. Another modified protocol was described by Wang et al.:[26] after fixing, staining solution employs 5% v/v acetic acid, 45% v/v ethanol and 0.125% (w/v) CBB R-250. Indeed, colloidal CBB-G staining is more sensitive than CBB-R in solvent solutions (8 ng).[14,18] The procedure requires multiple steps. Moreover, the need for two destaining solutions (first 5% v/v acetic acid, 40% v/v ethanol; then 3% v/v acetic acid, 30% v/v ethanol) renders the method prone to dye loss also by the stained proteins. Pink et al. [27] substituted acetic acid with phosphoric acid during the fixation (10%) and staining of the gels (8%) and also 5% aluminum sulfate (Al2(SO4)3) 14–18 hydrate in the staining solution. Al2(SO4)3 is used in the fabric industry as a mordant; however, Pink et al. stated that it also acts as an ion-bridging reagent, as Al3+ binds the dodecyl sulfate part of sodium dodecyl sulphate (SDS), by coagulation. Destaining is carried out in 2% phosphoric acid and 10% ethanol; afterwards gels are put in water overnight. Pink et al. modified this approach using a solution containing 10% ethanol, 0.02% of CBB G-250, and 8% phosphoric acid and 5% aluminum sulfate

colloidal CBB staining 0.02% CBB-G250 in 8% (w/v) phosphoric acid, 5% aluminum sulfate and 10% ethanol; for 3 h, after fixation 30% ethanol, 2% phosphoric acid; (C) CBB staining according to Neuhoff (gel was stained in 2% phosphoric acid, 10% ammonium sulfate, 20% methanol, 0.1% CBB G-250 overnight). CBB: Coomassie brilliant blue.

CBB R-250 SOLUTIONS

1. (0.1% (w/v) Coomassie brilliant blue (R-250), 25% (v/v) isopropanol, 10% (v/v) (glacial) acetic acid and 65% (deionized water)

2. (0.1% CBB R-250/ 40% MeOH/ 10% Glacial Acetic Acid) (Vf = 500 ml)

Dissolve CBB R-250 in 200 ml Methanol.
Add 250 ml dH2O.
Slowly add 50 ml Glacial Acetic Acid.
Filter solution through a Whatman No. 1 filter to remove any impurities.
Store at RT in a dark bottle (light sensitive).

DESTAIN SOLUTION

(10% MeOH/ 7% Glacial Acetic Acid) (Vf = 500 ml) To 415 ml dH2O add:
50 ml Methanol.
35 ml Glacial Acetic Acid (add slowly).
Stir to mix.
Store at RT.

GEL STORAGE SOLUTION

(5% Glacial Acetic Acid)(Vf = 500 ml) To 475 ml dH2O add:
25 ml Glacial Acetic Acid.
Stir to mix.
Store at RT.

De-Stain sol

for Coomassie - 1 L

20% methanol, 10% acetic acid

Reagents needed:

200 ml methanol or 400 ml ethanol
100 ml glacial acetic acid
700 ml ddH2O

Directions:

1) Add 100 ml of glacial acetic acid to 700 ml of ddH2O.

2) Add 200 ml of methanol or or 400 ml ethanol and mix.

3) Store at room temperature in a sealable container.

Staining Sol:

0.1% Coomassie blue, 10% acetic acid, 40% ethanol.

Add: 0.25 g Coomassie Blue R-250 100 mL ethanol 100 mL water

Stir until dye is completely dissolved, about one hour. Add 25 mL acetic acid and make to 250 mL with water. Store at room temperature in a dark bottle.