SDS-PAGE AND STAINING METHODS
Arvind Singh, Sushila Kala
Biomedical Sciences Department,
B. U., Jhansi, India
Gel Electrophoresis
Electrophoresis is the separation of charged particles through a solution or gel, under the influence of an electric field. Gel electrophoresis refers to the technique in which molecules are forced across a span of gel. Separation of macromolecules depends on 2 forces: charge and mass. During electrophoresis, macromolecules are forced to move through the pores when the electrical current is applied. Their rate of migration through the electric field depends on the strength of the field, size, and shape of the molecules, relative hydrophobicity of the samples, and on the ionic strength and temperature of the buffer in which the molecules are moving. After staining, the separated macromolecules in each lane can be seen in a series of bands spread from one end of the gel to the other.
Poly Acrylamide Gel Electrophoresis (PAGE)
The PAGE technique was introduced by Raymond and weintraub (1959). Polyacrylamide gels are formed from the polymerization of two compounds, acrylamide and N,N -methylene- bis-acrylamide. Bis is a cross-linking agent for the gels. The polymerization is initiated by the addition of ammonium persulfate along with either -dimethyl amino-propionitrile (DMAP) or N,N,N ,N ,- tetramethylethylenediamine (TEMED). The gels are neutral, hydrophillic, thermo-stable, transparent, strong, relatively chemically inert three-dimensional networks of long hydrocarbons crosslinked by methylene groups. Poly acrylamide gel can also withstand high voltage gradients, feasible to various staining and destaining procedures and can be digested to extract separated fractions or dried for autoradiography and permanent recording. PAGE separates protein molecules according to both size and charge. Polyacrylamide gel may be prepared so as to provide a wide variety of electrophoretic conditions. The pore size of the gel may be varied to produce different molecular sieving effects for separating proteins of different sizes. Proteins with molecular weights ranging from 10,000 to 1,000,000 may be separated with 7 to 12% acrylamide gels, while proteins with higher molecular weights require lower acrylamide gel concentrations. Conversely, gels up to 30% have been used to separate small polypeptides. The higher the gel concentration, the smaller the pore size of the gel and the better it will be able to separate smaller molecules. The percent gel to use depends on the molecular weight of the protein to be separated. Use 5% gels for proteins ranging from 60,000 to 200,000 daltons, 10% gels for a range of 16,000 to 70,000 daltons and 15% gels for a range of 12,000 to 45,000 daltons. There are two layers of gel, namely stacking or spacer gel, and resolving or separating gel.
Stacking gel:
The stacking gel is a large pore polyacrylamide gel (4%). This gel is prepared with Tris/HCl buffer pH 6.8 of about 2 pH units lower than that of electrophoresis buffer (Tris/Glycine). These conditions provide an environment for Kohlrausch reactions determining molar conductivity, as a result, SDS-coated proteins are concentrated to several fold and a thin starting zone of the order of 19 μm is achieved in a few minutes. This gel is cast over the resolving gel. The height of the stacking gel region is always maintained more than double the height and the volume of the sample to be applied.
Resolving gel:
The resolving gel is a small pore polyacrylamide gel (3 - 30% acrylamide monomer) typically made using a pH 8.8 Tris/HCl buffer. In the resolving gel, macromolecules separate according to their size. Resolving gels have an optimal range of separation that is based on the percent of monomer present in the polymerization reaction; for example an 8%, 10% and 12% resolving gel can effectively used for separating proteins between, 24 – 205 kDa, 14-205 kDa, and 14-66 kDa proteins, respectively. PAGE can be done under both native and denaturing conditions.
1. Under native conditions. (Blue Native PAGE, Clear Native PAGE)
2. Under denaturing conditions. (SDS-PAGE)
We will concentrate on denaturing polyacrylamide gel electrophoresis in the presence of sodium dodecylsulfate (SDS-PAGE) and a reducing agent.
The solution of proteins to be analyzed is first mixed with SDS, an anionic detergent which denatures secondary and non–disulfide–linked tertiary structures, and applies a negative charge to each protein in proportion to its mass. Without SDS, different proteins with similar molecular weights would migrate differently due to differences in mass charge ratio, as each protein has an isoelectric point and molecular weight particular to its primary structure. This is known as Native PAGE. Adding SDS solves this problem, as it binds to and unfolds the protein, giving a near uniform negative charge along the length of the polypeptide. SDS binds in a ratio of approximately 1.4 g SDS per 1.0 g protein (although binding ratios can vary from 1.1-2.2 g SDS/g protein), giving an approximately uniform mass:charge ratio for most proteins, so that the distance of migration through the gel can be assumed to be directly related to only the size of the protein. A tracking dye may be added to the protein solution to allow the experimenter to track the progress of the protein solution through the gel during the electrophoretic run.
Composition of stock solutions for use in SDS_PAGE:
1. Acrylamide / bis Acrylamide ( 30% T, 2.67% C)
(gm. Acrylamide + gm. bis-Acrylamide) x100
(Total volume)
gm. bis-Acrylamide x100
(gm.Acrylamide + gm. bis-Acrylamide)
Acrylamide 29.2 g / 100 ml.
N’ N’ bis-methylene – Acrylamide 0.8 g / 100 ml
Filtere and store at 4 0C in the dark (30 days maximum)
2. 1. 5 M Tris-HCl , pH 8. 8
18.15 g Tris base / 100 ml. (pH is adjusted with 6 N HCl and final volume was made to 100 ml with deionized water)
3. 0. 5 M Tris-HCl, pH 6. 8
6 g Tris base / 100 ml. (pH is adjusted with 6 N HCl and final volume was made to 100 ml with deionized water)
4. 10 % SDS
5. 4X sample buffer
1 M Tris –HCl, pH 6.8 2.5 ml
SDS 0.8 g
2-Mercapto ethanol 1.0 ml
Glycerol 3.0 ml
Bromophenol Blue 2.0 mg
Total volume 10 ml
6. 5X Running Buffer, pH 8. 3
Tris base 15g / lit.
Glycine 72 g / lit.
SDS 5 g / lit.
7. Separating Gel Preparation – 0.375 M Tris, pH 8. 8
S.No Reagents 10% 12% 15%
1:30% polyacrylamide (mL) 3.33 4 5
2:1.5 M Tris (pH 8.8) (mL)2.5 2.5 2.5
3:10% Ammonium persulfate(µL)100 100 100
4 10% SDS (µL) 100 100 100
5 TEMED (µl) 4 4 4
6 H20 (mL) 3.96 3.3 2.3
Total Volume (mL) 10 10 10
4 % Stacking Gel Preparation – 0.125 M Tris, pH 6.8
Deionized water 3.05 ml
1.5 M Tris-HCl, pH 8.8 1.25 ml
10 % SDS 50 µl
Acrylamide / Bis (30% stock) 0.665 ml
10% Ammonium per sulfate 25 µl
(Freshly prepared)
TEMED 5 µl
Total volume 5 ml
Safety and Practical Points
Acrylamide and bis-acrylamide are toxic as long as they are not polymerized
Procedure:
1. Clean glass plates and spacers. Set up gel plates with spacers and Place in casting stand.
2. Prepare resolving gel. Combine all but APS and TEMED. Fresh APS = 0.25 g APS + 0.85 ml dH2O.
3. Seal plates: Combine 200 µl resolving gel mix with 1 µl TEMED + 1 µl APS. Pour 100 µl down inside of each spacer.
4. Allow to polymerize at bottom (1-2 minutes). Add APS and TEMED to resolving gel mix (amounts can be increased 25%). Swirl gently.
5. Pour to ~5 mm below bottom of wells (~3.5 ml). Overlay with dH2O-saturated butanol. Will polymerize in 15-20 minutes. Pour off unpolymerized resolving gel mix. Wash with dH2O several times (squirt bottle). Place on side to drain dH2O completely.
6. Prepare stacking gel. Use APS made above. Pour 0.9 ml stacker. Introduce clean, dry comb at an angle to avoid trapping bubbles.
7. Make 1 L running buffer. Clamp gel to electrode stand. Place in tank and add buffer. Remove comb slowly. Wash out wells with reservoir buffer using syringe.
8. Load 5-10 µl of sample on bottom of well. Use lanes on outer edges last.
Running conditions:
At constant ampere of 16 mA till the dye reaches separating gel, then increased to 20 mA.
Staining of PAGE gels:
1. Coomassie Blue staining
Stain ˝ hr with 0.1% Coomassie blue R-250 in fixative (40% Methanol and 10 % glacial Acetic acid).
Destaining: Destain with several changes of 40% methanol and 10% glacial acetic acid to remove background.
2. Silver staining:
Silver staining is less compatible with the mass spectrometric analysis. Therefore, we don t recommend using silver staining for any samples that will subsequently be submitted for MS analysis. If you can not visualize bands with coomasie and you are not able to scale up your isolation of protein to reach coomasie stainable levels, you must use a mass spec compatible silver stain protocol.
• You must only use methanol and acetic acid during the fixing step.
• You must not use any solutions containing formaldehyde or glutaraldehyde to fix the gel.
• You should clean a glass or plastic tray with detergent and rinse it thoroughly.
• You should wear clean, disposable gloves and never touch the gel bands directly
Fixation
1) Fix gel in 150 ml 50% methanol + 5% acetic acid for 20 min.
2) Wash in 150 ml 50% methanol for 10 min.
3) Wash in water for 10 min.
(For 1 gel, mix 75 ml methanol, 7.5 ml acetic acid, and 67.5 ml water.)
Sensitizing
4) Incubate with 150 ml 0,02% sodium thiosulfate for 1 min.
5) Rinse with water for 1 min. 2X.
(0.02% Sodium Thiosulfate: For 1 gel, add 30mg sodium thiosulfate-5 hydrate to 150ml water.)
Silver reaction
6) Submerge gel in 150ml 0.1% silver nitrate with 0.08% formalin (37%) for 20 min.
7) Rinse with water for 1 min. 2X.
(0.1% Silver Nitrate and 0.08% Formalin (37%): For 1 gel, add 150 mg silver nitrate and 120ul 37% formalin to 150ml water.)
Developing:
Must make fresh solution
8) Incubate with 150ml 2% sodium carbonate with 0.04% formalin (37%) until desired intensity of staining occurs. If developer turns yellow, which it often does within 30 sec, then discard and replace with fresh 150ml developer.
(2% sodium carbonate + 0.04% Formalin: For 1 gel, add 6g sodium carbonate to 300ml water.
Just prior to use, add 120ul 37% Formaldehyde to the 300ml.)
Stopping
9) Wash gel in 150ml 5% acetic acid for 10 min.
(5% acetic acid: For 1 gel, add 7.5 ml acetic acid to 150ml water.)
Washing
10) Wash gel in water for 5 min.
Permanent Storage
11) Incubate gel in 150ml preserving solution for 20min.
(Preserving solution: For 1 gel, add 13.2ml glycerol (100% w/w) to 150 ml water.)
3. CuCl2 Staining:
This protocol is 2-3 times more sensitive than coomasie blue staining. It is much quicker, and the gels can be stored at 4° for many months without protein degradation.
Cupric Chloride Stain
0.3 M cupric chloride (5.1 g cupric chloride up to 100 ml with milliQ water)
Procedure
1. Run a standard SDS-PAGE gel.
2. Following electrophoresis, transfer directly to the Cupric Chloride Stain solution and shake for 5 minutes.
3. Rinse in milli Q water and transfer to saran wrap on a black background to visualize the bands.
4. The gels can be photographed under white light and stored for many months at 4°. The bands will not diffuse unless they are cut from the gel.
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