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An Introduction to Gel Electrophoresis

Gel electrophoresis is one of the most common tools used in a laboratory to separate and visualise macromolecules such as proteins, DNA or RNA. The main types of gels used during electrophoresis are agarose gels and polyacrylamide gels. Both of these gel types will be discussed, starting with agarose gel electrophoresis.

Agarose gel electrophoresis

What goes into the gel?

Agarose gels are typically used to analyse DNA and RNA. These gels, as the name suggests, contain agarose, a polysaccharide found in agar. Agar is extracted from various red algae, such as species of the genus Gracilaria, and is composed of agarose and agaropectin. Agar on its own can be used to produce agar plates (Petri dishes with agar containing nutrients for bacterial growth), yet agarose is mostly reserved for gels due to its physical and chemical properties that make it ideal for electrophoresis.

Agarose is usually available in powdered form and is dissolved in a buffer to make the gel. The amount of agarose used to make the gel defines the concentration of agarose in the gel, which is denoted as a percentage. Most gels have an agarose percentage of 0.7% to 2%. A higher percentage of agarose allows for more accurate separation of DNA or RNA samples that are small or very close in size. EDTA, SYBR Safe and Ethidium bromide and are other components that go into making the gel.

Gel preparation

The agarose is dissolved in buffers such as TAE or TBA; these buffers are called gel casting buffers. Buffers for electrophoresis contain ethylenediaminetetraacetic acid (shortened to EDTA), a chelating agent. A chelating agent inhibits DNA or RNA degradation by nucleases that require metal ions as co-factors. In this way, EDTA ensures that nucleic acid samples are more likely to remain intact.

A flask containing the gel solution is swirled and microwaved until all the agarose has dissolved. The glass will get hot so thick heat-protective gloves are used to take out the flask. The flask is swirled once again to ensure that the gel has a regular concentration of agarose throughout.

Once the agarose solution has cooled down, ethidium bromide or SYBR Safe is added to the solution. Ethidium bromide intercalates between base pairs of double-stranded DNA and fluoresces under UV light allowing samples to be visualised. However, ethidium bromide doesn’t stain RNA or single-stranded DNA as well as double-stranded DNA and can be mutagenic. SYBR Safe is a less mutagenic alternative.

Next, the solution is poured into a tray with rubber walls on two opposite ends to cool and solidify. A comb is carefully inserted into the cooling agarose solution to create holes that will form the sample loading ‘wells’.

Once the gel has cooled and solidified, the rubber walls are taken off and the gel is transferred to an electrophoresis tank. Running buffer is poured into the sides of the tank until the gel is submerged.

Before the sample are loaded onto the gel, loading buffer is added to them. The loading buffer is used for several purposes:

  1. It makes the sample denser than the TAE/TBE buffer, so the sample sinks to the bottom of the well without spilling outside.

  2. It contains a dye so the bands can be visualised and tracked as they move down the gel.

In a well adjacent to the samples, a “DNA ladder” is loaded. This contains various fragments of linear nucleic acids with known lengths, so the length of linear DNA/RNA samples can be estimated. The lid is placed on the tank and electrodes are attached to the power supply. The gel is now ready to run, and the current is turned on. The end of the tank away from the wells has the anode and will become positively charged while the top with the cathode, closer to the wells, will become negatively charged. Bands of the sample will migrate down the gel.

Before the dye in the loading buffer reaches the end of the gel, the current is turned off to prevent the sample from running off the gel. The gel is then visualised under UV light to identify the bands more clearly.

To run RNA on a gel, the samples must be protected from RNases, which are extremely abundant in the environment. A bleach gel (agarose gel with low levels of bleach) has been proposed as a method to overcome this. Alternatively, using equipment that is RNase-free as well as RNase inhibitors or inactivators can reduce RNase contamination risks. Urea could also be used to denature RNases. However, both the bleach gel and urea will also remove any RNA secondary structures.

Protocols (and references for the overview in this article) can be found in this list:

Some video demonstrations can be found in the following links:

Figure 1 shows a simple diagram of the stages involved in producing an agarose gel.

Figure 1: Stages of agarose gel production. Agarose chains extracted from red algae are used to produce a gel solution, which is poured into a gel casting tray. A comb produces wells in the gel as it solidifies. Note that rubber ends must be put on the tray before pouring. See Footnote 1 for image credits.

How does the gel work?

When agarose is dissolved in buffer, the agarose molecules becomes disordered. As the solution cools, the agarose chains aggregate to form helical fibres due to intermolecular hydrogen bonding. These fibres create a net-like structure that has regular pore sizes. Nucleic acids are net negatively charged molecules and travel towards the anode through these pores. The larger the sample of DNA or RNA, the longer it takes for it to “sieve” through the pores and migrate down the gel. Consequently, nucleic acids of smaller sizes migrate through the gel faster.

Plasmid DNA can be supercoiled and these types of DNA molecules are more compact. A supercoiled piece of DNA of a certain length would travel through the agarose gel faster than linear DNA of the same length. When interpreting the bands alongside the ladder, one must consider if a sample may contain supercoiled plasmids, nicked plasmids or linear DNA as the conformation of plasmid DNA will affect migration rate.

After nucleic acid bands are separated, further analysis can be carried out following gel extraction.

Polyacrylamide gel electrophoresis (PAGE)

What goes into the gel?

A polyacrylamide gel functions similarly to an agarose gel, but a net of cross-linked polyacrylamide chains provides the pores for the sample to migrate through. PAGE is often used for the separation of proteins. Solutions for PAGE gels include acrylamide, bisacrylamide, buffer, water, ammonium persulfate (APS) and tetramethylethylenediamine (TEMED).

Denaturing PAGE (or SDS-PAGE) gels for proteins use sodium dodecyl sulfate (SDS). SDS is a denaturing agent that disrupts intermolecular and intramolecular non-covalent interactions. SDS is an anionic substance and will coat proteins. This confers a negative charge to the polypeptides so samples will migrate down a gel towards the anode. SDS must be used alongside a reducing agent (β-mercaptoethanol) to remove any disulfide bridges within the tertiary structure.

Gels that attempt to conserve the structure and biological activity of the protein samples for further analysis are known as native-PAGE. There are several types of native-PAGE and these will be explained in the “How does the gel work?” section.

PAGE usually uses discontinuous buffer systems, which use buffers with varying pH and buffer ions for the gel and the running buffer.

Gel preparation

For SDS-PAGE, all the components described in the previous section are added sequentially, with TEMED added last. The solution is poured into the casting chamber and is left to polymerise. This forms the “resolving” gel. The resolving gel buffer is adjusted to a pH of around 8.8 using hydrochloric acid. Butan-2-ol can be added to the top of the gel to remove oxygen bubbles, as oxygen can diminish polymerisation. Once the gel is ready, the top of the resolving gel is rinsed with water and the “stacking” gel is produced. The stacking gel is prepared similarly to the resolving gel, but the buffer used has a lower pH of around 6.8. A comb is inserted into the stacking gel as the polyacrylamide polymerises to create the sample loading wells. Once the stacking gel has polymerised, running buffer is poured onto the top of this gel into a container that sits underneath the resolving gel then the comb is removed. The solvent for the samples contains β-mercaptoethanol, APS, TEMED, buffer, glycerol and bromophenol blue. Glycerol ensures the sample sinks to the bottom of the well and bromophenol blue, a marker dye, keeps track of the migrating protein. Native PAGE gels do not require reducing agents or denaturing agents.

How does the gel work?

Acrylamide, when in solution, has the capacity to polymerise. Monomers of acrylamide bond covalently to form long polyacrylamide molecules of various lengths. The bisacrylamide crosslinks these polyacrylamide molecules and creates a 3D net-like structure with pores that allow the sample to travel through. To initiate and speed up the process of polymerisation, ammonium APS and TEMED are added. Figure 2 shows a simple diagram of cross-linked polyacrylamide. A higher percentage of acrylamide will result in smaller pores and is ideal for separating smaller proteins.

Figure 2: Diagram showing the chemical structures of acrylamide, polyacrylamide, bisacrylamide and a section of crosslinked polyacrylamide chains. The repeating acrylamide unit in the cross-linked structure is denoted by the yellow brackets with the incomplete bonds signifying a bond to the repeating unit. Do bear in mind that the bonds lengths and angles were constructed for clarity and are not accurate.

Denaturing PAGE gels will separate based on the molecular weight of a protein. This is because:

  1. Once a protein has been fully denatured, its tertiary structure no longer influences how fast it moves through the pores in the cross-linked polyacrylamide.

  2. A protein’s native charge has a negligible impact on the rate of migration through the gel. SDS binds proteins regularly (around 1.4g of SDS/1g protein). Since SDS is negatively charged, the overall charge on the protein will become roughly proportional to its length and intrinsic charge of the polypeptide is masked.

There are several types of commonly used native PAGE. These include:

  • Clear native PAGE. This technique does not artificially charge samples and therefore separates based on native charge, size and shape.

  • Native blue PAGE. This is used to separate proteins by native molecular weights as the Coomasie Brilliant Blue R-250 dye is used to coat proteins with a negative charge, similarly to SDS. However, such a method will help retain tertiary structures of proteins and complexes, unlike SDS-PAGE.

PAGE often uses Tris-Glycine/Tris-HCl discontinuous buffer systems. There are different types of buffer systems that may be used as described in the following links:

Advantages specific to certain types of polyacrylamide gels can be partially attributed to different buffer systems. What remains consistent is the need for a “trailing ion” and a “leading ion”.

A discontinuous buffer system is used to concentrate the protein sample or “sharpen” the band. As the sample sits in a well, some polypeptides will be closer to the bottom of the well. These proteins enter the gel and start running before other polypeptides. To overcome this, the stacking gel ensures the proteins enter the resolving gel at the same time. Using the example of Tris-Glycine/Tris-HCl system, the stacking gel has a pH of 6.8 where glycine is mostly zwitterionic and chloride is negatively charged. Here, glycine has the lowest electrophoretic mobility and travels towards the anode the slowest; it acts as the trailing ion. Chloride travels the fastest as the leading ion while the protein has mobility between that of chloride and glycine. As chloride ions migrate rapidly down the gel and glycine trails behind, the proteins in between them become more concentrated and the band gets narrower. At a pH of 8.8 in the resolving gel, glycine becomes negatively charged and has higher electrophoretic mobility than the protein sample and migrates ahead of the concentrated band. The proteins are then free to separate. The trailing ion, leading ion and the pHs required for stacking vary between systems.

For native PAGE, the pH of the buffers must be carefully considered to maintain the physiological charge of the protein while also allowing stacking to occur.

After running the gel, protein bands can be visualised by staining. A commonly used staining agent is Coomassie Brilliant Blue R-250. As the Coomassie dye is anionic, staining an SDS-PAGE gel requires large amounts of dye to be added due to the interference with SDS that is already bound to the protein. Alternatively, the proteins can be analysed further with techniques such as Western blotting.

PAGE can also be done with DNA or RNA, with many materials and methods for the experiment changed to suit nucleic acids. There are some overlaps with the explained procedure for agarose gel electrophoresis. An example protocol can be found here.

Various protocols, summaries and learning resources for PAGE (also references for this article) can be found by clicking on the following links:

The following links contain videos on SDS-PAGE:

Other variations

There are many other forms of gel electrophoresis, such as capillary electrophoresis, pulsed-field gel electrophoresis, SDD-AGE and 2D gel electrophoresis. 2D gels separate samples by isoelectric point in 1 dimension and by molecular weight in the other giving a “map” of protein spots in a gel. This gives better separation of proteins in a sample for extraction and further analysis. Additionally, it gives 2 dimensions of information to identify each protein band.

2D gel electrophoresis can be used to study differences in protein produced in various tissue or cell types. A specific application of this involves characterising proteomic differences between healthy and cancer cells, where a proteome describes the assortment of proteins in an organism or tissue.

In summary, there are many different types of gel electrophoresis methods, and gel electrophoresis is a powerful analytical technique employed in many scientific experiments.


Kenta Renard

BSc Biochemistry

Imperial College London


  1. “Stages of agarose gel production” by Kenta Renard includes “File: Agarose Gel, with Comb inserted, in a Gel Tray (Front, angled view) - SketchUp.png” by Jacopo Werther which is licensed under CC BY-SA 4.0 and a cartoon adaptation of “2% agarose gel in a gel tray (front, angled)” by Joseph Elsbernd which is licensed under CC BY 2.0. “Stages of agarose gel production” by Kenta Renard is licensed under CC BY-SA 4.0.


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