Sunday, 26 October 2014

ELECTROPHORESIS: Principle, SDS PAGE, use and preparation of buffer, data analysis, gel preparation ( key chemicals used), schematic representation, separation of proteins by SDS PAGE, staining of protein gel, silver staining technique and documentation.



ELECTROPHOREESIS


CONTENT:


Principle, SDS PAGE, use and preparation of buffer, data analysis, gel preparation (key chemicals used), schematic representation.

Electrophoresis is the migration of charged molecules in response to an electric field. Their rate of migration depends on the strength of the field; on the net charge, size and shape of the molecules, and also on the ionic strength, viscosity, and temperature of the medium in which the molecules are moving. As an analytical tool, electrophoresis is simple, rapid, and highly sensitive. It is used analytically to study the properties of a single charged species, and as a separation technique. 

There are a variety of electrophoretic techniques, which yield different information and have different uses. Generally, the samples are run in a support matrix, the most commonly used being agarose and polyacrylamide. These are porous gels, and under appropriate conditions, they provide a means of separating molecules by size. We will focus on those methods used for proteins. These can be denaturing or non-denaturing. Nondenaturing methods allow recovery of active proteins and can be used to analyze enzyme activity or any other analysis that requires a native protein structure. Two commonly used techniques in biochemistry are sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) and isoelectric focusing (IEF). SDS-PAGE separates proteins according to molecular weight and IEF separates according to isoelectric point. This laboratory exercise will introduce you to SDS-PAGE.

SDS-PAGE


The gel matrix used is a cross linked acrylamide polymer. This electrophoretic method separates the proteins according to size (and not charge) due to the presence of SDS. The dodecyl sulfate ions bind to the peptide backbone, both denaturing the proteins and giving them a uniform negative charge. The gels we will be running use a discontinuous system, meaning that they have 2 parts. One is the separating gel, which has a high concentration of acrylamide and acts as a molecular sieve to separate the proteins according to size. Before reaching this gel, the proteins migrate through a stacking gel, which serves to compress the proteins into a narrow band so they all enter the separating gel at about the same time. The narrow starting band increases the resolution. This part of the gel has a lower concentration of acrylamide to avoid a sieving effect. The stacking effect is due to the glycine in the buffer, the low pH in the stacking gel, and the higher pH in the running buffer. At the low pH, the glycine has little negative charge, and thus moves slowly. The chloride ions move quickly and a localized voltage gradient develops between the 2. As the gel runs, the low pH of the stacking gel buffer is replaced by the higher pH in the running buffer. This maintains a discontinuity in the pH and keeps the glycine moving forward (any glycine molecules behind would acquire a higher charge and speed up). Since there is no real sieving going on, the proteins (which have intermediate mobility) form a tight band, in order of size, between the slower glycine and the faster chloride ions. The separating gel buffer has a higher pH, so the glycine molecules become more negatively charged and move past the proteins, and the voltage gradient becomes uniform. The proteins slowdown in the smaller pore size of the separating gel and separate according to size. Exercise: You will be given protein molecular weight standards, several different solutions containing individual proteins, and a sample of the same serum you used in the protein quantitation lab. Your job is to determine the molecular weights of the individual proteins and the major components in the serum sample. You will run each sample on 2 gels, one you prepare yourself and a commercial precast gel, and compare the results. Before doing electrophoresis, you must know the amount of protein in each sample. Determine the protein concentrations of each of your samples using a protein assay before coming to the lab to do any electrophoresis. For this exercise, the only sample of unknown protein concentration is the serum that you used for one of your unknowns last week. The amount of protein to be loaded depends on the thickness and length of the gel, and the staining system to be used. Using the Coomassie Blue staining system, as little as 0.1 mg can be detected, but more will be easier to see. As a guide, use 0.5–5 mg for pure samples (one or very few proteins) and 20–60 mg for complex mixtures where the protein will be distributed amongst many protein bands. Overloading will decrease the resolution. Protocol: The apparatuses used in gel casting or running electrophoresis vary; make sure you look over the appropriate manuals before you operate. Caution: Unpolymerized Acrylamide is a Neurotoxin. Be Careful! Do not pour unpolymerized acrylamide down the sink, wait for it to polymerize and dispose of it in the trash. TEMED (N, N, N', N'-tetramethylethelenediamine) is also not very good for you and is very smelly; avoid breathing it. Open the bottle only as long as necessary, or use it in the hood. 

1. Make sure gel plates are clean and dry. Do not get your fingerprints on them or the acrylamide will not polymerize properly.
 2. Prepare gel solutions (separating and stacking), but do not add polymerizing agents, APS and TEMED (this would start the polymerization).
 3. Lay the comb on the unnotched plate and mark (on the outside, using a Sharpie) about 1 cm below the bottom of the teeth. This will be the level of the separating gel. If available, use an alumina (opaque, white) plate, for the notched plate, as this conducts heat away from the gel more efficiently than glass. Set up the gel plates, spacers, and plastic pouch in the gel casting as described in the manufacturer’s directions. When everything is completely ready, add TEMED to the separating gel solution, mix well, and pour it between the plates, up to the mark. Wear gloves if you pour directly from the beaker. You can also use a disposable pipette. Work quickly or the solution will polymerize too soon. Carefully layer isopropanol (or water-saturated butanol) on top of acrylamide so it will polymerize with a flat top surface (i.e., no meniscus). Do this at the side and avoid large drops, so as not to disturb the gel surface. When the leftover acrylamide in the beaker is polymerized, the acrylamide between the plates will also be ready.
4. If you are running the gel on the same day, prepare samples while the acrylamide is polymerizing. Otherwise, wait until you are ready to run the gel.
(i) You will need a sample of each unknown substance, plus the molecular weight standards. Prepare samples in screw-cap microcentrifuge tubes. The protein content should be at 1–50 mg in 20–30 mL sample. The total sample volume that can be loaded depends on the thickness of the gel and the diameter of the comb teeth. For Genei apparatuses, this is ~ 30 mL/well. To prepare the sample, mix 7–10 mL of the sample (depending on protein concentration) +20 mL 2X sample buffer containing 10% b-mercaptoethanol (BME). Use the BME in the hood - it stinks! For dilute samples, mix 40 mL of the sample and 10 mL 5X sample buffer and add 2 mL of BME. Heat to 90°C for 3 minutes to completely denature proteins. It is important to heat samples immediately after the addition of the sample buffer. Partially denatured proteins are much more susceptible to proteolysis and proteases are not the first proteins to get denatured. (Heat samples to 37°C to redissolve SDS before running the gel if samples have been stored after preparation).
(ii) If you want the proteins in the sample to retain disulfide bonds, do not add BME. If both reduced and nonreduced samples will be run on the same gel, leave at least 3–4 empty wells between samples, since the BME will diffuse between wells and reduce proteins in adjacent samples.
(iii) MW Stds: 7 mL of Rainbow stds +10 mL of sample buffer (do not make in advance). Heat to 37°C before use.
 5. After the separating gel has polymerized, drain off the isopropanol. Add TEMED to the stacking gel solution, pour the solution between the plates, and insert the comb to make wells for loading samples. The person putting in the comb should wear gloves. Keep an eye on this while it’s polymerizing and add more gel solution if the level falls (as it usually does), or the wells will be too small.
 6. after polymerization, do not cut the bag; we reuse them. The gel may be stored at this point by taping the bag shut to prevent drying. When ready to run the gel: mark the position of each well, since they are difficult to see when full.
7. Remove comb and rinse wells with running buffer. See the manual directions for setting up the gels in the buffer chambers. The apparatus can run 2 gels simultaneously. There is a blank plate to use when running only one. Fill the upper chamber with running buffer first and check for leaks. Adjust the plates if necessary. Load the samples using a micropipettor with gel-loading tips (these are longer and thinner than the normal tips). This will be demonstrated. Do not load samples in the end wells. Make sure to write down which sample was loaded in each well.
8. Electrophoresis (takes 1–2 hours). Connect the gel apparatus to the power supply and run at 15 mA/gel until the tracking dye (blue) moves past the end of the stacking gel. Increase the current to 20–25 mA/gel but make sure the voltage does not get above 210 V. Run until the blue tracking dye moves to the bottom of the separating gel. For the BioRad apparatus do not exceed 30 mA, regardless of the number of gels. 9. Disassemble the apparatus and carefully separate the gel plates using a flat spatula. Cut off the stacking gel and any gel below the blue tracking dye. Note the color of each of the molecular weight standards, as they will all be blue after staining. Wash 3X with distilled water. Place the gel in a plastic staining container and add Coomassie Blue staining solution. Keep it in this 1 hour overnight. Wash again with water. You can wrap the gel in plastic wrap and Xerox or scan it to have a copy. The gel may also be dried.

Data Analysis

Measure the length of the gel (since you cut off the bottom, this is the distance traveled by the dye). Measure the distance traveled by each of the molecular weight standards. Measure the distances of each unknown band. For samples lanes with many bands (serum in this exercise), measure all bands in those with just a few and the major bands in those that have many. Prepare a standard curve by plotting log MW versus relative mobility (Rf, distance traveled by protein divided by distance traveled by dye). Use this and the mobility of bands from your fractions to determine the MW of the unknown proteins. (Review standard curves from the protein quantitation lab if necessary.) MW of proteins that do not run very far into the gel or run near the dye front will not be accurate. If you have reduced and unreduced samples, compare the number of bands and MW of each to determine the number of subunits.

Gel Solutions
1.     Separating gel: (15 mL, enough for two gels) 10% acrylamide. 40% Acrylamide/bisacrylamide mix 3.55 mL. 1.5 M tris pH 8.8, 3.75 mL, H2O 7.4 mL, 10% SDS 150 mL, 10% ammonium persulfate (APS) 150 mL (prepared fresh), TEMED 6 mL.
2.     2. Stacking gel: (5 mL) 5% acrylamide. Compresses the protein sample into a narrow band for better resolution. 40% Acrylamide/bisacrylamide mix 0.625 mL. 0.5 M tris pH 6.8, 1.25 mL, H2O 3.0 mL, 10% SDS 50 mL, 10% APS 50 mL, TEMED 5 mL.
3.     3. 2X sample buffer (10 mL)—store in the freezer for an extended time.
4.     4. SDS must be at room temperature to dissolve.
5.     5. H2O 1.5 mL, 0.5 M Tris pH 6.8, 2.5 mL, 10% SDS (optional) 4.0 mL, glycerol 2.0 mL, BPB 0.01%, b-mercaptoethanol (optional) 0.1 mL.
6.     6. Running buffer (5L) 30 g Tris Base, 144 g glycine, dissolve in sufficient H2O to make 1.5 L and put into final container. Add 1.5 g SDS (Caution: do not inhale dust). When adding SDS, avoid making too much foam, which makes measuring and pouring difficult. Final pH should be around 8.3, but do not adjust it or the ionic strength will be too high and the gel will not run properly. If the pH is way off, it was made incorrectly or is old and has some contamination. The running buffer can also be made more concentrated (5X or 10X) and diluted as needed to save bottle space.
Electrophoresis is defined as the separation (migration) of charged particles through a solution or gel, under the influence of an electrical field. The rate of movement of particle depends on the following factors.
1. The charge of the particle.
2. Applied electric field.
3. Temperature.
4. Nature of the suspended medium.

What is Gel Electrophoresis?

Gel electrophoresis is a method that separates macromolecules—either nucleic acids or proteins—on the basis of size, electric charge, and other physical properties. A gel is a colloid in a solid form. The term electrophoresis describes the migration of charged particles under the influence of an electric field. “Electro” refers to the energy of electricity. “Phoresis,” from the Greek verb phoros, means “to carry across.” Thus, gel electrophoresis refers to the technique in which molecules are forced across a span of gel, motivated by an electrical current. Activated electrodes at either end of the gel provide the driving force. A molecule’s properties determine how rapidly an electric field can move the molecule through a gelatinous medium. Many important biological molecules such as amino acids, peptides, proteins, nucleotides, and nucleic acids, possess ionizable groups and, therefore, at any given pH, exist in solution as electrically charged species, either as cations (+) or anions (–). Depending on the nature of the net charge, the charged particles will migrate to either the cathode or the anode.

How does this Technique Work?

Gel electrophoresis is a technique used for the separation of nucleic acids and proteins. Separation of large (macro) molecules depends upon 2 forces: charge and mass. When a biological sample, such as proteins or DNA, is mixed in a buffer solution and applied to a gel, these 2 forces act together. The electrical current from one electrode repels the molecules, while the other electrode simultaneously attracts the molecules. The frictional force of the gel material acts as a “molecular sieve,” separating the molecules by size. 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.

Agarose

There are 2 basic types of materials used to make gels: agarose and polyacrylamide. Agarose is a natural colloid extracted from seaweed. It is very fragile and easily destroyed by handling. Agarose gels have very large “pore” size and are used primarily to separate very large molecules, with a molecular mass greater than 200 kdal. Agarose gels can be processed faster than polyacrylamide gels, but their resolution is inferior. That is, the bands formed in the agarose gels are fuzzy and spread far apart. This is a result of pore size and cannot be controlled. Agarose is a linear polysaccharide (average molecular mass about 12,000) made up of the basic repeat unit agarobiose, which composes alternating units of galactose and 3, 6-anhydrogalactose. Agarose is usually used at concentrations between 1% and 3%. Agarose gels are formed by suspending dry agarose in an aqueous buffer, then boiling the mixture until a clear solution forms. This is poured and allowed to cool to room temperature to form a rigid gel.
Polyacrylamide

There are 2 basic types of materials used to make gels: agarose and polyacrylamide. The polyacrylamide gel electrophoresis (PAGE) technique was introduced by Raymond and Weintraub (1959). Polyacrylamide is the same material that is used for skin electrodes and in soft contact lenses. 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. In this way, the percentage of polyacrylamide can be controlled in a given gel. By controlling the percentage (from 3% to 30%), precise pore sizes can be obtained, usually from 5 to 2000 kdal. This is the ideal range for gene sequencing, protein, polypeptide, and enzyme analysis. Polyacrylamide gels can be cast in a single percentage or with varying gradients. Gradient gels provide a continuous decrease in pore size from the top to the bottom of the gel, resulting in thin bands. Because of this banding effect, detailed genetic and molecular analysis can be performed on gradient polyacrylamide gels. Polyacrylamide gels offer greater flexibility and more sharply defined banding than agarose gels.
Mobility of a molecule = (applied voltage) × (net charge of the molecule)/ friction                                                                                                                                                                                             
                                                of the molecule (in the electrical field)
V (velocity) = E (voltage) × q (charge)/f (frictional coefficient).


Polyacrylamide Gel Electrophoresis

Polyacrylamide is the solid support for electrophoresis when polypeptides, RNA, or DNA fragments are analyzed. Acrylamide plus N, N’-methylene-bis-acrylamide in a given percentage and ratio are polymerized in the presence of ammonium persulfate and TEMED (N, N, N’, N’-tetra-methyl-ethylene-diamine) as catalysts.
Safety and Practical Points

Acrylamide and bis-acrylamide are toxic as long as they are not polymerized.
Buffer (usually Tris) and other ingredients (detergents) are mixed with acrylamide before polymerization.
Degassing of acrylamide solution is necessary before pouring the gel because O2 is a strong inhibitor of the polymerization reaction.
Polyacrylamide Gel Electrophoresis of Proteins

Under nondenaturing conditions.
Under denaturing conditions.
Isoelectric focusing.
These techniques are used to analyze certain properties of a protein such as: isoelectric point, composition of a protein fraction or complex, purity of a protein fraction, and size of a protein. We will concentrate on denaturing polyacrylamide gel electrophoresis in the presence of sodium dodecyl sulfate (SDS-PAGE) and a reducing agent (DTT, or dithioerithritol, DTE). The protein is denatured by boiling in “sample buffer,” which contains:
1. Buffer pH 6.8 (Tris-HCl).
2. SDS.
3. Glycerol.
 4. DTT or DTE.
5. Bromophenol blue (tracking dye).

Discontinuous Polyacrylamide Gel Electrophoresis
This type of polyacrylamide gel consists of 2 parts:
1.     The larger running (resolving) gel
2.     The shorter upper stacking gel.
The running gel has a higher percentage (usually 10%–15%) of acrylamide and a Tris-HCl buffer of pH 8.8.
The stacking gel usually contains 5% acrylamide and a Tris-HCl buffer of pH 6.8.
The buffer used in SDS-PAGE is Tris-glycine with a pH of about 8.3.

Determination of the Molecular Weight of a Polypeptide by SDS-PAGE

Since all polypeptides are wrapped with SDS and thus are strongly negatively charged, they migrate through the running gel according to their size (small polypeptides migrate faster than large ones!). There is a linear relationship between the log of the molecular weight of the polypeptide and its migration during SDS-PAGE. Standard polypeptides have to be run on the same gel and a curve of their migration versus the log of their molecular weight has to be generated.

PREPARATION OF SDS-POLYACRYLAMIDE GELS
Materials
·        Casting gel unit for electrophoresis
·        Siliconized Pasteur pipettes
·        Syringes equipped with blunt stub-nosed needles
·        Vacuum chamber for degassing gels
·        Micropipettes
·        (10–300 mL) Stock 30%T:0.8%C acrylamide monomer
·        1.5 M Tris-HCl buffer, pH 8.8
·        10% (w/v) SDS
·        10% (w/v) ammonium persulfate
TEMED acrylamide is a powerful neurotoxin. Do not breathe powder or otherwise come in contact with the monomer. Wear gloves at all times.
Separation gel mixed just prior to use
• 20 mL of acrylamide monomer
• 15 mL of Tris-HCl Buffer, pH 8.8
• 0.6 mL of 10% (w/v) SDS
• 24.1 mL of H2O.
 Stacking gel mixed just prior to use
• 2.66 mL of acrylamide monomer
• 5.0 mL of Tris buffer, pH 8.8
• 0.2 mL of 10% (w/v) SDS
• 12.2 mL of H2O


Procedure

1. Assemble your slab gel unit with the glass sandwich set in the casting mode with 1.5-mm spacers in place.
2. Prepare a separating gel from the ingredients listed.
3. Add the separating gel to a side arm flask, stopper the flask, and attach to a vacuum pump equipped with a cold trap. Turn on the vacuum and degas the solution for approximately 10 minutes. During this period, gently swirl the solution in the flask.
4. Turn off the vacuum, open the flask, and add 200 mL of ammonium persulfate and 20 mL of TEMED to the solution.
5. Add the stopper to the flask and degas for an additional 2 minutes while gently swirling the solution to mix the 2 accelerators. Use this solution within a few minutes of mixing, or it will gel in the flask.
6. Transfer the degassed acrylamide solution to the casting chamber with a Pasteur pipette. Gently fill the center of the glass chamber with the solution by allowing the solution to run down the side of one of the spacers. Be careful not to introduce air bubbles during this step.
7. Adjust the level of the gel in the chamber by inserting a syringe equipped with a 22-gauge needle into the chamber and removing excess gel.
8. Immediately water layers the gels to prevent formation of a curved meniscus. Using a second syringe and needle, add approximately 0.5 mL of water to the chamber by placing the tip of the needle at an angle to a spacer and gently allowing the water to flow down the edge of the spacer and over the gel. Add an additional 0.5 mL of water to the chamber by layering it against the spacer on the opposite side of the chamber. Done appropriately, the water will form a layer over the gel, and a clear line of demarcation will be observed as the gel polymerizes.
9. After 30 minutes, the gel should be polymerized. If degassing was insufficient, or the ammonium persulfate not fresh, the polymerization may take an hour or more. When the gel is polymerized, lift the gel in its casting chamber and tilt to decant the water layer.
10. Prepare a stacking gel from the listed ingredients.
11. Degas the stacking gel as in step 3.
12. Add 75 mL of ammonium persulfate and 10 mL of TEMED to the stacking gel and degas for an additional 2 minutes.
13. Add approximately 1 mL of stacking gel to the gel chamber and gently rock back and forth to wash the surface of the separating gel. Pour off the still-liquid stacking gel and dispose of properly. Remember that liquid acrylamide is extremely hazardous!
14. Add fresh stacking gel until it nearly fills the chamber, but allow room for the insertion of a Teflon comb used to form sample wells. Carefully insert a Teflon comb into the chamber. Adjust the volume of the stacking gel as needed to completely fill the spaces in the comb. Be careful not to trap any air bubbles beneath the combs. Oxygen inhibits polymerization, and will subsequently result in poor protein separations.
15. Allow the gels to polymerize for at least 30 minutes prior to use.

SEPARATION OF PROTEIN STANDARDS: SDS-PAGE
Materials
·        10% SDS-polyacrylamide gel
·        Protein standards
·        2X-SDS sample buffer
·        1X-SDS electrophoresis running buffer (Tris-Glycine + SDS)
·        0.001% (w/v) bromophenol blue Micropipettes with flat tips for electrophoresis wells.
Procedure
1. Remove the Teflon combs from the prepared gels by gently lifting the combs from the chamber. Rinse the wells (formed by the removal of the combs) with distilled water and drain it off.
2. Fill the wells and the chamber with running buffer.
3. Prepare aliquots of a known protein standard by mixing equal parts of the protein standard with 2X sample buffer.
4. Using a micropipette, add the sample to the bottom of a well. Add the blue to a separate well.
5. Remove the gel from its casting stand and assemble it into the appropriate slab unit for running the electrophoresis. Be sure to follow the manufacturer’s directions for assembly.
6. Pour a sufficient quantity of running buffer into both the lower and upper chambers of the electrophoresis apparatus until the bottom of the gel is immersed in buffer, and the top is covered, while the electrodes reach into the buffer of the upper chamber. Be careful not to disturb the samples in the wells when adding buffer to the upper chamber.
7. Assemble the top of the electrophoresis apparatus and connect the system to an appropriate power source. Be sure that the cathode (+) is connected to the upper buffer chamber.
8. Turn on the power supply and run the gel at 20 mA constant current per 1.5 mm of gel. For example, if 2 gels are run, each with 1.5-mm spacers, the current should be adjusted to 40 mA. One gel with 1.5-mm spacers should be run at 20 mA, while a gel with 0.75-mm spacers should be run at 10 mA.
9. When the tracking dye reaches the separating gel layer, increase the current to 30 mA per 1.5-mm gel.
10. Continue applying the current until the tracking dye reaches the bottom of the separating gel layer (approximately 4 hours).
11. Turn off and disconnect the power supply. Disassemble the gel apparatus and remove the glass sandwich containing the gel. Place the sandwich flat on paper towels and carefully remove the clamps from the sandwich.
12. Working on one side of the sandwich, carefully slide 1 of the spacers out from between the 2 glass plates. Using the spacer or a plastic wedge as a lever, gently pry the glass plates apart without damaging the gel contained within.
13. Lift the bottom glass plate with the gel and transfer the gel to an appropriate container filled with buffer, stain, or preservative. The gel may at this point be used for Coomasie Blue staining, silver staining, enzyme detection, Western blots, or more advanced procedures, such as electroblotting or electro elution. If prestained protein standards were used, the gels may be scanned directly for analysis. Place the gel into 50% methanol and gently rock the container for about 30 minutes prior to scanning. This can be accomplished by placing the gels into a flat dish and gently lifting the edge of the disk once every 30 seconds. There are commercially available rocker units for this purpose. If the gel is to be dried, use a commercial gel dryer such as (SE 1160 Slab Gel Dryer). Following the manufacturer’s directions demonstrates a dried and stained gel containing a series of proteins of known molecular weights.
14. Plot the relative mobility of each protein against the log of its molecular weight. Relative mobility is the term used for the ratio of the distance the protein has moved from its point of origin (the beginning of the separating gel) relative to the distance the tracking dye has moved (the gel front). The ratio is abbreviated as Rf. Molecular weight is expressed in daltons, and presents a plot of the relative molecular weight of protein standards against the log of their molecular weight.

COOMASSIE BLUE STAINING OF PROTEIN GELS
·        Protein gel 
·        0.25% (w/v) Coomassie Brilliant Blue R 250 in methanol-water-glacial acetic acid (5/5/1), filtered immediately before use
·        7% (v/v) acetic acid
·        Commercial destaining unit (optional)
Procedure
1. Place a gel (prepared as in Exercise 2) in at least 10 volumes of Coomassie Blue staining solution for 2–4 hours. Rock gently to distribute the dye evenly over the gel.
2. At the conclusion of the staining, wash the gels with water a few times.
3. Place the gels into a solution of 7% acetic acid for at least 1 hour.
4. If the background is still deeply stained at the end of the hour, move the gels to fresh 7% acetic acid as often as necessary. If a commercial destainer is available, this will decrease the time required for stain removal. Follow the manufacturer’s directions for use of the destainer.
5. Place the gels into containers filled with 7% acetic acid as a final fixative.
6. Photograph the gels or analyze the gels spectrophotometrically.
Notes
Coomassie Brilliant Blue R 250 is the most commonly used staining procedure for the detection of proteins. It is the method of choice if SDS is used in the electrophoresis of proteins, and is sensitive for a range of 0.5 to 20 micrograms of protein. Within this range, it also follows the Beer-Lambert law and, thus, can be quantitative as well as qualitative. The major drawback is the length of time for the procedure and the requirement for destaining. Over staining results in a significant retention of stain within the gel, and thus, a high background stain, which might obliterate the bands. The length of time for staining must be carefully monitored, and can range from 20 minutes to several hours. If maximum sensitivity is desired, one should try 2 hours for a 5% gel and 4 hours for a 10% gel. Destaining must be monitored visually and adjusted accordingly.

SILVER STAINING OF GELS
Materials
·        Protein gel from
·        45% (v/v) methanol + 12% (w/v) acetic acid.
·        5% (v/v) methanol + 7% (w/v) acetic acid.
·        10% Glutaraldehyde
·        0.01 M Dithiothreitol
·        Silver nitrate solution Sodium citrate/formaldehyde
·        Kodak Farmer’s Reducer or Kodak Rapid Fixer

Procedure
1. Fix gels by gently rocking them in a solution of 45% methanol/12% acetic acid until the gels are completely submerged. Fix for 30 minutes at room temperature. 2. Remove the fixative and wash twice for 15 minutes each with 5% ethanol/ 7% acetic acid. (Gels thicker than 1 mm require longer washing.)
3. Soak the gels for 30 minutes in 10% glutaraldehyde.
 4. Wash thrice with deionized water, 10 minutes each.
5. Place in dithiothreitol for 30 minutes.
6. Place in silver nitrate solution for 30 minutes.
7. Wash for 1 minute with deionized water. Dispose of used silver nitrate solution immediately with continuous flushing. This solution is potentially explosive when crystals form upon drying.
8. Place in sodium citrate/formaldehyde solution for 1 minute.
9. Replace the sodium carbonate/formaldehyde solution with a fresh batch, place gels on a light box, and observe the development of the bands. Continue to rock gently as the gel develops.
10. When the desired degree of banding is observed (and before the entire gel turns black), withdraw the citrate/formaldehyde solution and immediately add 1% glacial acetic acid for 5 minutes.
11. Replace the glacial acetic acid with Farmer’s reducer or Kodak Rapid Fixer for 1 minute. Remove Farmer’s reducer and wash with several changes of deionized water.
12. Photograph or scan the gel with a densitometer, which produces typical silver stained protein gel.
13. For storage, soak the gel in 3% glycerol for 5 minutes and dry between dialysis membranes under reduced pressure at 80–82°C for 3 hours. Alternatively, place the wet gel into a plastic container (a storage bag will do) and store at room temperature. If desired, the gels may be dried between Whatman 3-MM filter paper for autoradiography, or dried using a commercial gel dryer.

DOCUMENTATION
Materials
·        Polaroid camera (Fotodyne Foto/Phoresis I or equivalent) or  35-mm camera equipped with macro lens
·        Stained gel

Procedure
1. Photograph the gels.
2. Use the photographs or negatives to measure the distance from the point of protein application (or for 2 gel systems, the line separating the stacking and separating gels) to the final location of the tracking dye near the bottom of the gel.
3. Measure the distance from the point of origin to the center of each band appearing on the gel.
4. Divide each of the values obtained in step 3 by that obtained in step 2 to obtain the relative mobility (the Rf value) for each band.
5. Using either the graph of Rf values and molecular weights from Exercise 2, compute the molecular weights of each band.
Optional
Scan the negative with a densitometer and compute Rf values based on the distances from the point of origin to the peak tracing for each protein band. Integration of the area of each peak will yield quantitative data, as well as the molecular weight.

SCHEME OF ELECTROPHORESIS





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