Electrophoretic Mobility Shift Assay (EMSA)


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The electrophoretic mobility shift assay (EMSA) is a biochemical procedure used to elucidate binding between proteins and nucleic acids. In this assay a radiolabeled nucleic acid and test protein are mixed. Binding is determined via gel electrophoresis which separates components based on mass, charge, and conformation.

This video shows the concepts of EMSA and a general procedure, including gel and protein preparation, binding, electrophoresis, and detection. Applications covered in this video include the analysis of chromatin-remodeling enzymes, a modified EMSA that incorporates biontinylation, and the study of binding sites of bacterial response regulators.

EMSA, the electrophoretic mobility shift assay, also known as the gel shift assay, is a versatile and sensitive biochemical procedure. EMSA elucidates binding between proteins and nucleic acids by detecting a shift in bands in gel electrophoresis.

This video describes the principles of EMSA, provides a general procedure, and discusses some applications.

DNA replication, transcription, and repair, as well as RNA processing are all critical biochemical processes. They all involve binding between proteins and nucleic acids. Many serious diseases and disorders are associated with modifications in this binding. EMSA is a technique for qualitatively determining whether a specific protein binds to a specific nucleic acid. First, the nucleic acid is labeled, usually with radioactive phosphorus-32, to create a probe. Then the test protein and nucleic acid probe are mixed. When a protein binds to a nucleic acid probe, the resulting complex has greater mass and a different conformation than the nucleic acid alone.

Once bound, the complexes are analyzed with gel electrophoresis. In this technique, an electric field forces macromolecules to migrate through a gel matrix. The components separate based on mass, charge, and conformation. Electrophoresis can separate protein-DNA complexes from unbound probes. Since they have different masses and conformations, they will migrate through the gel at different rates and separate. The separation is easily detected, thanks to the presence of the radioactive phosphorus, and proves the protein successfully binds to the given nucleic acid. To verify the identification of the protein, a "supershift assay" uses an antibody with a known affinity to the protein. This has the added benefit of further shifting the complex, increasing resolution.

Now that we've seen the principles, let's see it in the lab.

To begin the procedure, the protein must be isolated. To do this, molecular biology techniques are used to express the protein in cells, and then purify.

The nucleic acid is amplified and labeled to create a probe. Labeling is done through incubation for 10 min with dCTP containing radioactive phosphorus-32. A radiation-safe workbench and protective equipment are required.

The gel is then prepared. The gel needs to be non-denaturing, to prevent the protein from altering conformation and potentially unbinding from the probe during electrophoresis. Polyacrylamide gels have pore sizes of 5 to 20 nm and are useful for short probes up to 100 base pairs in length. Agarose gels have pore sizes of 70-700 nm and are useful for larger probes.

With the protein and nucleic acid probe now prepared, we proceed to binding. A TRIS buffer solution is prepared and the protein and probe are added. The pH should be similar to physiological conditions, and salt concentration sufficient to prevent the protein from forming weak bonds with non-target nucleic acids. The reaction proceeds for 20-30 min at 4 °C.

The next step is electrophoresis. A buffer of low ionic strength and a pH similar to that used in the binding reaction is utilized. It yields a "caging effect" that stabilizes complexes, increases mobility, and reduces heat generation. After electrophoresis, the components of the gel are transferred onto filter paper. In a dark room, the filter paper is then exposed to film. If the protein binds to the probe, two distinct labeled regions will be visible in the transfer. One representing the complex, and a separate one representing the unbound probe. The separation demonstrates that the protein successfully bound to the nucleic acid.

Now that we've seen the basic procedure, let's look at some applications.

Chromatin is the tightly packed complex of DNA and proteins found eukaryotic cells. Chromatin-remodeling enzymes modify the structure to open the DNA to transcription. As this changes the mobility of the complex, EMSA can be used to explore the binding activity of the enzyme.

An alternate approach to labeling takes advantage of the interaction between DNA and the methyltransferase enzyme. A cofactor can be modified to bind permanently to DNA via methyltransferase. Rather than labeling with phosphorus-32, the cofactor is conjugated to biotin, which is advantageous because it's not radioactive. Because the cofactor is site-specific in its attachment, it’s relevant to genotyping, methylation detection, and gene delivery. The biotinilated nucleic acids are detected through ultraviolet fluorescence.

When environmental stimuli activate histidine kinases, a “response regulator” is phosphorylated. This in turn binds to DNA, affecting transcription, which can be studied by EMSA. For instance, a response regulator in Desulfovibrio vulgaris was demonstrated to bind to the gene of interest. EMSA was used to verify that the binding took place.

You've just watched JoVE's video on the electrophoretic mobility shift assay. You should now understand its principles of operation, the steps in its procedure, and its major operating parameters.

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JoVE Science Education Database. Biochemistry. Electrophoretic Mobility Shift Assay (EMSA). JoVE, Cambridge, MA, (2020).

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