Each protein has a specific molecular weight. This property is used by scientists to separate them based on their sizes.
SDS-PAGE is a technique to separate proteins using an electric current, solely based on their sizes, that is, by their molecular weights. This separation occurs through a technique involving electrophoresis, and it is run on a polyacrylamide gel.
To better understand, let’s look at little closer at each of these terms.
SDS stands for Sodium Dodecyl Sulfate, a detergent.
PAGE is the abbreviation of Poly Acrylamide Gel Electrophoresis. And this is where we get the combined term SDS-PAGE.
Electrophoresis is a procedure that relies on an electric current to separate macromolecules, specifically in this case, proteins in a mixture. The separation here is solely based on the protein’s molecular weights.
Polyacrylamide, is a polymer that the gel is made up of.
Researchers use this technique for many reasons and in many ways. For example, from out of the entire protein mixture in the sample, you might want to send a specific protein of a certain known size for downstream analysis such as mass spectroscopy.
In this article we will also understand why SDS-PAGE is such a useful technique for biochemists, as well as explore the basic concept of this protein separation technique, and the utility of SDS in this procedure.
In this article:
In electrophoresis, under the influence of an electric field, charged molecules travel through a medium towards a specific electrode.
The negatively charged particles travel to the positive electrode (anode), and positively charged particles toward the negative electrode (cathode).
If all molecules in a mixture of proteins are similarly charged, then during electrophoresis, the constituents are separated based on their sizes.
All proteins in the mixture will travel towards the same electrode given that all proteins in the mixture have similar charge, but the smallest protein will migrate the farthest, and the largest protein will travel the shortest distance towards that electrode.
So, the underlying working principle of protein electrophoresis is very similar to that of DNA electrophoresis.
However, a fundamental difference remains between DNA and proteins, which makes their electrophoresis procedures different.
DNA are negatively charged molecules due to their sugar-phosphate backbone. When electrophoresed, they automatically migrate towards the positive electrode.
In contrast, a protein molecule may carry an overall positive or negative charge or may be uncharged depending on its amino acid sequence.
Further, amino acids, due to having both NH2+ and COO- groups and consequently the protein molecule, may have variable overall charge as the pH varies.
So, if you think about it, when a mixture of proteins, at a certain pH, are put in an electric field for electrophoresis, depending on their overall charge, some of the protein molecules may migrate towards the cathode, others towards the anode, and some may not migrate at all.
To overcome this uncertainty, and to ensure that all proteins have an overall similar charge and migrate towards one electrode, a special treatment is done to the protein mixture before electrophoresis.
The protein sample mix is treated with the detergent SDS (Sodium Dodecyl Sulfate). In the next section we’ll take a deeper look at why this is done.
The detergent Sodium Dodecyl Sulfate (SDS) is an integral part of most protein electrophoresis procedures. Its role is so important that it is part of the name of the procedure – SDS-PAGE.
The underlying objective of SDS-PAGE is to separate proteins only on the basis of their molecular weights. Any other property such as shape and intrinsic charge should not interfere.
SDS is a negatively charged molecule. It confers an overall negative charge to all protein molecules when added to the sample.
Figure 1. Depicts SDS molecules uniformly covering a linearized polypeptide with a negative charge. By making proteins in a mixture negatively charged, all proteins are attracted to the positive electrode.
As a result, the intrinsic overall charges of the proteins, due to their amino acid behavior at the experimental pH, are nullified, And all proteins in the sample become negatively charged.
This ensures that during electrophoresis all constituent protein molecules in the sample will migrate towards the positive electrode, and the separation will be solely based on size (molecular weight).
SDS facilitates unfolding the secondary and tertiary structures of proteins in the sample mix.
In order to separate proteins only based on their molecular weights, the protein molecules need to be denatured to their linear forms. SDS greatly facilitates unfolding the secondary and tertiary structures of proteins in the sample mix.
The SDS molecule has a hydrophobic region as well as an ionic part. This hydrophobic region interacts with and unfolds hydrophobic regions of proteins.
Figure 2. Shows the hydrophobic (purple) and ionic (blue) regions of SDS.
The ionic part of SDS disrupts non-covalent interactions within proteins. This results in denaturing the protein molecules to their primary structure.
SDS helps to denature the proteins in the sample. Also, SDS ensures that all proteins become negatively charged and travel towards the positive electrode so that separation of the proteins is solely based on molecular weight.
But a few other treatments of your protein sample are still required for a successful SDS-PAGE procedure.
In this section we will take a look at these treatments, and understand why they are done.
The three-dimensional shape of a protein molecule is largely governed by hydrogen bonds – think about alpha helix and beta sheets.
For SDS-PAGE, since protein samples need to be linearized into their primary structures, these hydrogen bonds need to be broken.
For this, as part of preparation for SDS-PAGE, the sample is heated for a few minutes at around 95°C. Heating destroys hydrogen bonds (figure 3).
Figure 3: Proteins are denatured to their linear primary structure by treatment with heat prior to electrophoresis
Treatment with BME and DTT
The protein sample, as part of its preparation for electrophoresis, is treated with BME (Beta Mercapto Ethanol) and/ or DTT (Dithiothreitol).
These compounds are chemically categorized as “reducing thiols.”
The role of sulfur-containing amino acids – like cysteine, in determining protein tertiary structure is that they form disulfide bridges.
Treating the protein sample with BME or DTT break these disulfide bridges, facilitating the denaturation of sample proteins to their unfolded state (figure 4).
Figure 4. DTT or BME disrupt the disulfide bridges between sulfur containing amino acids like cysteine and help to linearize the polypeptide backbone
In electrophoresis, consider the gel as a molecular sieve, a tightly woven mesh. Just like an ordinary sieve – say a coffee filter, it allows smaller particles to pass through easily, while the larger particles are left behind due to their bigger size.
In protein electrophoresis, all proteins in the sample, due to their negative charge (because of SDS treatment), migrate towards the positive electrode.
However, due to the sieve-like property of the gel, the proteins in the sample are separated based on size – the smallest protein migrate farthest towards the positive electrode while the largest ones do not migrate as far.
For a more detailed conceptual understanding of how a gel facilitates separation based on size please refer to this article on DNA gel electrophoresis.
PAGE is a polymer of acrylamide. Let us take a detailed look at the chemistry of this polymerization.
Bis-acrylamide serves as the cross-linking agent, linking up poly-acrylamide chains – this ultimately leads to the formation of a three-dimensional network of polyacrylamide chains that serves as the molecular mesh.
Since bis-acrylamide is only a cross-linking agent joining acrylamide polymer chains, the ratio of bis-acrylamide:acrylamide in the gel is about 1:35.
Roles of TEMED and APS in gel formation
Here is a chemical hierarchy that leads to the acrylamide polymerization reaction:
- A compound TEMED, when added to APS, drives formation of persulfate free radicals.
- In turn, these persulfate free radicals, when added to acrylamide, leads to the formation of acrylamide free radicals. They are reactive and polymerize with acrylamide monomers. This initiates acrylamide polymer formation. So as you can see in figure 5 below, acrylamide monomers do not readily react until activated by TEMED.
For the actual procedure of gel construction, acrylamide and bis-acrylamide are first mixed. TEMED and APS are then added which immediately initiate the gel formation reaction.
Figure 5. Chemistry behind polyacrylamide gel formation - the reaction between acrylamide and bis-acrylamide, and the roles played here by TEMED and APS.
As soon as TEMED and APS are added, the gel mix is poured between two glass plates where the gel solidifies. The apparatus with the two glass plates fixed together is technically called a gel cassette.
The process of making a gel is called casting. Figure 6 demonstrates the steps of casting a polyacrylamide gel.
Figure 6. The casting steps of a polyacrylamide gel.
Generally, for protein electrophoresis, the same polyacrylamide gel has two parts, a stacking part and a resolving gel.
A resolving gel solution (acrylamide ~10%, pH ~8.8) forms the bottom of the casting tray, and so it is poured before the stacking gel as shown in figure 6A.
Figure 6A. Resolving gel is being pipetted into the casting tray.
Isopropanol is then layered over the resolving gel before it solidifies (figure 6B). This prevents air bubbles and protects the resolving gel from drying out.
Also, this helps in flattening the top layer of the resolving gel rather than getting curvy.
Figure 6B. Isopropanol is layered over the resolving gel before it solidifies
The stacking gel, which has a lower acrylamide percentage (~4%), a different pH (~6.8) and ionic composition than the resolving portion, is then poured into the casting tray as illustrated in figure 6C.
Figure 6C. Stacking gel solution is being pipetted into the casting tray.
After pouring the stacking gel mix, a comb is placed on top to create “wells” (figure 6D).
Figure 6D. A comb is placed on top of the stacking gel to create wells.
Once the gel is ready, the protein samples are added into these wells for the electrophoretic separation to start.
Adding protein samples into the wells is technically called loading the gel.
Consider a race as an anology. At the first whistle, the candidates get ready, steady and on their mark. At the next and final whistle, the actual race starts.
The stacking gel serves as this first whistle. In the first few minutes of the electrophoresis, the racing candidates, that is, the proteins undergoing electrophoresis, get arranged and concentrated at the same level.
After this, the proteins enter the resolving gel, where the actual race starts – the smallest size protein is the winner.
Passing the proteins through the stacking gel before they reach the resolving gel ensures an optimal resolution (separation) of the proteins. So, during casting, the resolving gel is first cast. Once it solidifies, the stacking gel is cast on top of it.
The cast gel cassette is put into a running apparatus.
The running apparatus has electrodes connected to a source of electricity and is filled with a running buffer solution.
This buffer, due to its ions, facilitates the flow of electric current – the driving force behind electrophoretic separation.
Buffer solutions are critically important in biochemical experimentations, particularly with proteins.
They serve to maintain a certain pH; for example, around 6.8 and 8.8 for stacking and resolving gels respectively. And they conduct electricity.
Think about how the running buffer conducts electricity through the gel for the electrophoresis to take place
In SDS-PAGE three different buffers are used:
- for casting the gel
- for sample preparation and loading (please see SDS-PAGE loading buffer below)
- for running the gel
You may find our buffer handbook really helpful because it not only goes into detail about common biological buffers, but it also details buffer chemistry.
Loading samples into the wells of gel
We have already discussed the theory behind how protein samples need to be treated for SDS-PAGE separation – with SDS, BME and heat – to unfold them to their primary structure. Let us now take a brief glance at the procedural details.
An SDS-PAGE loading buffer is prepared. It contains SDS, BME and sometimes another disulfide bridge disruptor like DTT or BME, the chemicals we already discussed as necessary to denature the proteins in the sample.
Additionally, the loading buffer contains two more chemicals, glycerol and a dye, such as bromophenol blue(figure 7).
Glycerol in the loading buffer increases its density. This ensures that the sample drops to the bottom of the well during gel loading.
This prevents spillover and contamination of samples between the wells which may produce erroneous experimental results.
The dye bromophenol blue is added in the loading buffer for tracking purposes to monitor how far the samples have migrated in the gel.
Dyes like bromophenol blue ensure that electrophoresis is stopped at the proper time so that proper separation is obtained. Dyes also ensure that the samples do not run all the way out of the gel. If the samples run out of the gel, you won’t be able to analyze them.
Figure 7. Sample loading into the gel.
Bromophenol blue is negatively charged at the same pH at which SDS-PAGE is done. This ensures that the dye migrates towards the positive electrode with the SDS-covered protein samples.
One of the wells in the gel is loaded with a protein ladder.
The ladder is an assortment of multiple highly purified polypeptides of predetermined and varying sizes.
These polypeptides, due to their sizes differing between each another, migrate different known distances in the gel during electrophoresis.
As shown in the figure 8, this creates a band pattern akin to the rungs of a ladder – hence the name protein ladder. Each rung of the ladder corresponds to a polypeptide of a certain size (molecular weight).
Figure 8. Sample loading into the gel.
The protein ladder is used to estimate the sizes of polypeptides in the experimental sample.
Each band of the migrated ladder corresponds to a single molecular weight. By visually noting how far a polypeptide in the experimental sample has migrated as compared to the bands of the ladder, its molecular weight can be estimated.
Other minor utilities of a protein ladder are:
- Visual confirmation of how far a polypeptide of a certain size might have migrated at a certain time during electrophoresis while the gel is running.
- Confirmation of protein transfer to a membrane from the gel during subsequent procedures such as Western blot.
The general experimental approach is as follows:
- The gel is taken out of the electrophoresis chamber and washed with double distilled (dd) water to remove any running buffer.
- The gel is treated with a fixative solution (generally methanol-acetic acid based). The polypeptide bands get fixed in their respective positions in the gel matrix so that they do not diffuse and produce erroneous results.
- The gel is then treated with a staining solution that contains a dye. The stain binds to the protein bands in the gel so that they can be visually seen and interpreted.
- The gel is washed to remove excess stain. This step is technically called destaining. This is done to remove nonspecific background stain coloration, which improves sensitivity.
During staining, everything including the proteins and the gel matrix gets covered by the dye.
Since the entire gel is stained, the polypeptide bands do not stand out visually. This is why destaining is important. Destaining removes the dye from areas where it is non-specifically bound, like the gel matrix.
Only the polypeptides remain stained because they are specifically bound by the dye and can be visually recognized.
Gel staining facilitates quick visualization of the protein bands.
Some other experimental inferences can also be deduced after visualization of the polypeptide bands in the stained-destained gel.
For example, if there are differences in band number or intensity between lanes of the gel, that indicates a difference in protein abundance between samples respectively loaded in those wells.
The stained polypeptide bands may also be excised from the gel and sent out for downstream analysis such as mass spectroscopy.
While some gels are stained following SDS-PAGE, some are not. Instead, the unstained gel is used in downstream analytical procedures. In such cases, staining the gel is avoided because it may hamper the downstream experimental steps.
A very common protein analytical technique, where SDS-PAGE is done but the gel isn’t stained, but used for other downstream steps is Western blot.
- SDS-PAGE separates proteins based on their molecular weights.
- SDS is a detergent that confers an overall negative charge to all proteins in the sample. This ensures two important things. One, all proteins migrate towards the positive electrode. Secondly, protein separation is strictly and solely based on molecular weight.
- Other than SDS treatment, the sample is heated and is treated with BME or DTT. All of this is done to denature all proteins in the sample to their primary structure.
- The gel is made up of the polymer polyacrylamide. TEMED and APS aids in acrylamide and bisacrylamide polymerization.
- SDS-PAGE forms the stepping stone for two downstream protein-analytical methods – mass spectroscopy and Western blot. This highlights the importance of SDS-PAGE in modern research.
When you read or hear ground-breaking science about a new protein, remember, SDS-PAGE was a key experimental procedure behind all of that top-notch discovery.
- Wood EJ. 1993. Gel electrophoresis of proteins. Bios Scientific. doi.org/10.1016/0307-4412(93)90119-K
- Garfin DE. 1990. One dimensional gel electrophoresis. Methods Enzymol. 182: 425-441
- Peters TN. 1990. Gel electrophoresis of proteins. Oxford University Press. doi.org/10.1016/0307-4412(91)90147-Z