Introduction
SDS PAGE or Sodium Dodecyl Sulphate-Polyacrylamide Gel Electrophoresis is a technique used for the separation of proteins based on their molecular weight. It is a technique widely used in forensics, genetics, biotechnology and molecular biology to separate the protein molecules based on their electrophoretic mobility.
Principle of SDS-PAGE
The principle of SDS-PAGE states that a charged molecule migrates to the electrode with the opposite sign when placed in an electric field. The separation of the charged molecules depends upon the relative mobility of charged species.
The smaller molecules migrate faster due to less resistance during electrophoresis. The structure and the charge of the proteins also influence the rate of migration. Sodium dodecyl sulphate and polyacrylamide eliminate the influence of structure and charge of the proteins, and the proteins are separated based on the length of the polypeptide chain.
When proteins are separated by electrophoresis through a gel matrix, smaller proteins migrate faster due to less resistance from the gel matrix. Other influences on the rate of migration through the gel matrix include the structure and charge of the proteins.
In SDS-PAGE, the use of sodium dodecyl sulfate (SDS, also known as sodium lauryl sulfate) and polyacrylamide gel largely eliminates the influence of the structure and charge, and proteins are separated solely based on polypeptide chain length.
SDS is a detergent with a strong protein-denaturing effect and binds to the protein backbone at a constant molar ratio. In the presence of SDS and a reducing agent that cleaves disulfide bonds critical for proper folding, proteins unfold into linear chains with negative charge proportional to the polypeptide chain length.
Polymerized acrylamide (polyacrylamide) forms a mesh-like matrix suitable for the separation of proteins of typical size. The strength of the gel allows easy handling. Polyacrylamide gel electrophoresis of SDS-treated proteins allows researchers to separate proteins based on their length in an easy, inexpensive, and relatively accurate manner.
Role of SDS in SDS-PAGE
SDS is a detergent present in the SDS-PAGE sample buffer. SDS along with some reducing agents function to break the disulphide bonds of proteins disrupting the tertiary structure of proteins.
Ingredients
Most SDS PAGE sample buffers contain the following: SDS (sodium dodecyl sulphate, also called lauryl sulphate), b-mercaptoethanol (BME), bromophenol blue, glycerol, and Tris-glycine at pH 6.8. BME is added to prevent oxidation of cysteines and to break up disulfide bonds.
Bromophenyl blue is a dye that is useful for visualizing your sample in the well and tracking its progress through the gel. Glycerol is much more dense than water and is added to make the sample fall to the bottom of the sample well rather than just flow out and mix with all the buffer in the upper reservoir. The interesting components are the buffer and the SDS.
SDS is an ionic detergent that binds to the vast majority of proteins at a constant ratio of 1.4 gm SDS/gm protein. A few proteins like tubulin do not bind at this ratio and this is one reason why some proteins migrate anomalously (there are other reasons as well so you shouldn’t put too much faith in the apparent molecular weight estimated from an SDS PAGE gel).
Since SDS is an anionic detergent it imparts a negative charge to all the proteins in your sample. More importantly, these charges swamp the inherent charge of the proteins and give every protein the same charge-to-mass ratio. Because the proteins have the same charge-to-mass ratio, and because the gels have sieving properties, mobility becomes a function of molecular weight. But what about running gels, stacking gels, electrode buffer, and all these different pHs?
The velocity of a charged particle moving in an electric field is directly proportional to the field strength and the charge on the molecule and is inversely proportional to the size of the molecule and the viscosity of the medium. Adding a gel with sieving properties (that is a gel where the resistance to the motion of a particle increases with particle size) increases the differences in mobility between proteins of different molecular weights. This is the basis of separation. The problem now becomes how to line up all the proteins in an orderly fashion at the starting gate. That’s where the discontinuous pH part comes in.
Laemmli gels are composed of two different gels (stacker and running gel), each cast at a different pH. In addition, the gel buffer is at a third, different pH. The running gel is buffered with Tris by adjusting it to pH 8.8 with HCl. The stacking gel is also buffered with Tris but adjusted to pH 6.8 with HCl.
The sample buffer is also buffered to pH 6.8 with Tris HCl (note all the chloride ions – they will become important in a minute). The electrode buffer is also Tris, but here the pH is adjusted to a few tenths of a unit below the running gel (in this case 8.3) using only glycine – nothing else. We run our gels at constant voltage.
So here’s what happens when you turn on the power. Glycine is a weak acid and it can exist in either of two states, an uncharged zwitterion, or a charged glycinate anion (that is to say, negatively charged). At low pH it is protonated and thus uncharged.
At higher pH it is negatively charged. When the power goes on the glycine ions in the running buffer want to move away from the cathode (the negative electrode) so they head toward the sample and the stacking gel.
The pH there is low and so they lose a lot of their charge and slow down. Meanwhile, in the stacker and sample the highly mobile chloride ions (which are also negatively charged) move away from the cathode too.
This creates a narrow zone of very low conductance (in other words very high electrical resistance) in the top of the stacking gel. Because V=IR almost all of the voltage that you put across the gel (110 Volts is typical for stacking) is concentrated in this small zone. The very high field strength makes the negatively charged proteins move forward.
The trick, however, is that they can never outrun the chloride ions. If they did they would find themselves in a region of high conductance and very low field strength and would immediately slow down.
The result is that all the proteins move through the stacker in a tight band just behind the moving front of chloride ions. Behind them, the pokey glycine ions straggle along as best they can (they do move, but with lower mobility than the chloride ions).
The effect of this moving zone of high voltage is that all the proteins reach the running gel at virtually the same time so that migration of the proteins is truly a function of molecular size and not some complicated function of how carefully you loaded the gel and when you started the voltage.
When the big caravan of ions hits the running gel everything changes. The pH goes way up and the glycine becomes deprotonated (and thus more negatively charged). The mobility of the glycine goes way up and the mobility of the proteins goes way down (due to the sieving properties of the gel).
The result is that the glycine races past the protein and the proteins are no longer in a narrow zone of very high resistance (and very high electric field). They find themselves in a much more relaxed, uniform electric field where they can chill out a bit. Move at their own pace.
Tips for running good gels:
1. After pouring the running gel, carefully overlay it with ethanol or another imiscible liquid. This will give you a nice flat surface. Also, since polymerization of acrylimide is inhibited by oxygen it will speed up polymerization.
2. For the mini-gels we run the minimum protein loading per well (single band) is 0.1 µg for standard Coomassie staining and 2 ng for silver staining. I haven’t tested it but my impression is that Simply Blue staining is within a factor of two as sensitive as standard Coomassie staining.
3. The maximum protein loading per well (for a mixture of proteins of different sizes) is about 40 µg. If you exceed amount this your gel will look like crap.
4. KCl causes SDS to precipitate. If you samples contain KCl you should dilute them or methanol precipitate them and resuspend them in 1X sample buffer. With low concentrations of KCl (<200 mM) you can run them on the gel but you should loaed every lane with sample buffer containing the same concentration of KCl (even if they are blanks). This will help the gel run a little less anomalously.
5. If your sample buffer turns yellow, it is at the wrong pH.
Materials Required
1.Power Supplies: It is used to convert the AC current to DC current.
2.Gels: These are either prepared in the laboratory or precast gels are purchased from the market.
3.Electrophoresis Chambers: The chambers that can fit the SDS-PAGE gels should be used.
4.Protein Samples: The protein is diluted using SDS-PAGE sample buffer and boiled for 10 minutes. A reducing agent such as dithiothreitol or 2-mercaptoethanol is also added to reduce the disulfide linkages to prevent any tertiary protein folding.
5.Running Buffer: The protein samples loaded on the gel are run in SDS-PAGE running buffer.
6.Staining and Destaining Buffer: The gel is stained with Coomassie Stain Solution. The gel is then destained with the destaining solution. Protein bands are then visible under naked eyes.
7.Protein Ladder: A reference protein ladder is used to determine the location of the protein of interest, based on the molecular size.
Protocol of SDS-PAGE
1.Preparation of the Gel
All the reagents are combined, except TEMED, for the preparation of gel.
When the gel is ready to be poured, add TEMED.
The separating gel is poured in the casting chamber.
Add butanol before polymerization to remove the unwanted air bubbles present.
The comb is inserted in the spaces between the glass plate.
The polymerized gel is known as the “gel cassette”.
2.Sample Preparation
Boil some water in a beaker.
Add 2-mercaptoethanol to the sample buffer.
Place the buffer solution in microcentrifuge tubes and add protein sample to it.
Take MW markers in separate tubes.
Boil the samples for less than 5 minutes to completely denature the proteins.
3.Electrophoresis
The gel cassette is removed from the casting stand and placed in the electrode assembly.
The electrode assembly is fixed in the clamp stand.
1x electrophoresis buffer is poured in the opening of the casting frame to fill the wells of the gel.
Pipette 30ml of the denatured sample in the well.
The tank is then covered with a lid and the unit is connected to a power supply.
The sample is allowed to run at 30mA for about 1 hour.
The bands are then seen under UV light.
Applications of SDS-PAGE
The applications of SDS-PAGE are as follows:
1.It is used to measure the molecular weight of the molecules.
2.It is used to estimate the size of the protein.
3.Used in peptide mapping
4.It is used to compare the polypeptide composition of different structures.
5.It is used to estimate the purity of the proteins.
6.It is used in Western Blotting and protein ubiquitination.
7.It is used in HIV test to separate the HIV proteins.
8.Analyzing the size and number of polypeptide subunits.
9.To analyze post-translational modifications.
There are some limitations associated with the concept and applying the methods of the SDS page when it comes to identifying a particular strain on the basis of its DNA or RNA. Since the electrophoretic system helps us to visualise the protein molecules and the structures as it can identify the mass of the proteins, it is not helpful to identify the DNA or RNA structure of a strain. DNA and RNA are primarily composed of nucleic acids and when a component is contaminated with nucleic acid, the structure will not be visible on the SDS page gel.
In the system of SDS page, the SDS refers to an anionic detergent that is primarily used to denature or take away the basic characteristics of protein molecules in a sample. The SDS, which is negatively charged, will destroy the structures of protein molecules and then it will be strongly attracted toward the node of the electric field. This is how it helps us to separate the proteins depending on the mass.
The SDS page cannot determine the purity of the protein molecules. To test the purity, we have to perform various quantification methodologies like UV-Vis. Systems like electrophoresis are generally used to visualise the protein structure and separate the protein molecules on the basis of the mass or shape and structure. That is why pointing out the amount of impurity present in a protein molecule is not possible with the help of performing SDS-page or any other electrophoretic system.
