How to Select the Analysis Methods for Your Protein Project

Proteins can be analysed in a multitude of ways using a plethora of techniques. When it comes to protein purification, there are certain pieces of information about your protein that you are always interested in collecting. Any other aspects you may need to measure are decided by the nature of your research project.

For protein purification, the key pieces of information are identity, purity, size homogeneity, activity and concentration of your target protein (it is also worth trying to get some information about any key impurities as well).

Determining protein purity

Without doubt, the most common technique for determining protein purity is denaturing gel electrophoresis by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE). This technique separates proteins by size and allows various detection techniques to be used. Classic methods include Coomassie and silver staining, but more and more pre-labeling using a fluorescent dye is gaining in popularity. The classic staining methods use photographic detection using CCD digital imagers (or film if you're lab is old school :) which are robust but less sensitive and quantitative compared to fluorescent pre-labelling.

Alternatives to SDS-PAGE for purity analysis include 2-D PAGE, size exclusion chromatography, and mass spectrometry (Matrix-assisted laser desorption/ionization; MALDI-MS).

Measuring protein identity

The most common technique used for protein identity is western blotting. Western blotting uses denaturing SDS-PAGE gel electrophoresis followed by transfer of the separated proteins to a membrane. These proteins are then detected by a specific antibody and a secondary functionalized antibody which enables detection by chemiluminescence or fluorescence.

Mass spectrometry in combination with reverse phase chromatography, can provide an easy and fast complement or alternative to the antibody-based detection step in western blotting. The main drawback is that you need to have access to a mass spectrometer, a significant piece of kit and often a shared service. The main principle for confirming protein identity using mass-spectrometry is to trypsinate the SDS-PAGE gel band of interest prior to mass analysis. To identify the peptides, Electrospray Ionization (ESI) connected on-line with reverse phase chromatography is common. The mass-spectrum of peptide species after trypsinization provides a unique fingerprint for most proteins, which can be identified using a database lookup.

Use SEC in combination with SDS-PAGE for size homogeneity

Perhaps the most robust and powerful way of determining size homogeneity is to first separate your sample using size-exclusion chromatography (SEC), collect the fractions and then run a SDS-PAGE gel containing reducing agents such as dithiothreitol (DTT) on the fractions. The reducing agent breaks di-sulfide bridges between cysteine residues and the gel shows single-chain sub-units of the different sizes, if cysteins are causing multimerization. The textbook example is the combined purification and analysis of IgG as exemplified in the image above. The only drawback we can think of with this method is that if you have a low concentration of the protein in your sample, it may be difficult to detect. You also need to choose a SEC column with the right separation range for your protein of course.

Alternatives to this approach include using light-scattering or mass-spectrometry after the SEC step. Again, these detection techniques involve investment in expensive instruments.

Estimating concentration

As the subhead suggests, we think regardless of what measurement technique you use, you are likely to end up with nothing more than an estimation of the concentration. That said, measuring concentration is a chapter of its own (something we’re going to discuss in detail in future posts) but unfortunately no protein concentration assay method exists that is either specific to proteins or uniformly sensitive to all protein types (i.e. not affected by differences in protein composition). It is therefore important to choose the method that is most compatible with the sample and will give enough information for you to move forward with you research. 

For example, one of the most common methods when you want to check the expression level of you target protein from cultivation is to do a rough estimation with SDS-PAGE; it will show if you are on track. If you want to measure the total protein concentration the tried and tested methods are the Coomassie (Bradford) protein assay, BCA protein assay (also known as Smith assay) and UV absorbance at 280nm. 

Each of these methods has its own set of advantages and disadvantages. However, these methods give only an estimation of the total protein concentration. Because no method can be considered the ideal assay method for all circumstances, most researchers have more than one type of protein assay available in their laboratories. The BCA Protein Assay and Bradford Protein Assay methods are complementary and cover most samples, with both based on detection of color change. BCA is a two-step protocol including a Protein-copper chelation and secondary detection of the reduced copper. Bradford is a protein-dye binding and direct detection of the color change associated with the bound dye.

When choosing an assay somethings to consider:

•Compatibility with the sample type and components (e.g. in lysis buffer) that may interfere with the protein and/or the reagents in the assay used

•The concentration range of the assay and required sample volume. For example the Bradford assay works in a concentration range of 125–1,000 μg BSA /ml and the BCA assay in a working range of 20-2000μg BSA/ml

• Protein compositional differences which end up in different amount of color in the final solution and may give wrong concentrations- choose assay and protein standard which will minimise this error

•Speed and convenience for the number of samples to be tested

•Availability of spectrophotometer or plate reader

If your protein is an enzyme with activity in a specific enzymatic assay, using an assay may be an easy way to find out where in your eluted purification fractions the target protein is. You will then be able to detect your protein through all purification steps and have full control of the design of the purification protocol and quality of the obtained preparation. The activity is also an insurance that the protein is obtained in its native state.

We will go through the methods described in upcoming posts when we delve into our DHFR project. 

In the meantime, thanks for reading and if you have any questions let us know via the comments section below.  

Pairing your Protein with a Purification Tag

With the introduction of affinity tagging of proteins, protein purification was dramatically simplified; generic protocols could now be used, which enabled much more efficient and easy protein purification. This led to tags that do more than just purify protein, such as improving the solubility or stability.

In this post we’ll look at the characteristics, pros and cons of the most common tags used in protein purification today.

The “classic” His-tag

The histidine tag is by far the most commonly used tag for protein purification today. The reason for this is simple – it is so small that it is unlikely that it will interfere with the structure or function of the protein. This means that you don’t necessarily have to remove it before using the purified protein (one example when the his-tag often needs to be removed is for structure determination using X-ray crystallography). Another great benefit of using this tag is that purification is quite straightforward, and there is a great selection of ready-to-use purification products in a multitude of formats available to choose from. Different chromatography media are available that will provide different trade-offs between recovery, capacity and purity.

Adding histidine tag means that you typically add 4-6, sometimes up to 10, histidine residues to either the N- or the C-terminal of your protein. The aromatic group of the histidine residues bind to chelated di-valent metal ions. Nickel is the most commonly used, but Cobalt, Zinc and Copper can also be used. Zinc is the best choice for the environment. Regardless of the ion, imidazole is always used for the elution.

The main drawback of this tag is that it often requires some optimization of the imidazole concentration in your sample and in the buffers for column equilibration and wash in order to minimize binding of other host cell proteins with high histidine content.

Strep-tag™ II

Strep-tag II is a peptide tag that binds very specifically to Streptactin™, which is a modified version of streptavidin. Being small, it shares the benefits of the His-tag, and adds significant improvement in the purity you can expect.

In addition to the chromatography media being more expensive and having much lower binding capacity than media for purification of his-tagged proteins, the agent used for elution, desthiobiotin, is more expensive than imidazole.

GST for purity and solubility

Another very common tag is the enzyme Glutathione-S-Transferase (GST). It binds very specifically to glutathione immobilized on chromatography media, and therefore often gives very high purity. Another benefit is that it can also increase the solubility of the protein it is fused to. However, being big (26 kDa) it often needs to be cleaved off in order to eliminate interference with structure and function of your protein.

While there are chromatography media with high binding capacity, the kinetics of the binding is slow. The latter means that sample loading needs to be done at low flow-rates and therefore will take longer compared to e.g. a his-tagged protein.

MBP tag

Maltose Binding Protein (MBP) is another protein tag that can be used for purification and as an alternative to GST. Whilst providing the same benefit of high specificity and ability to improve solubility of your protein, it is larger than GST, so typically requires removal prior to using your protein. Lower binding capacity compared to GST and a more limited number of purification products available make this tag a second choice if the GST-tag for some reason does not work.

FLAG™ tag

If none of the to the tags discussed above work, the FLAG peptide-tag is a small tag that binds very specifically to a specific antibody currently only available on one type of chromatography media. In addition to the high specificity and thereby purity that can be expected, another benefit of this tag is that it is small, and therefore is unlikely to interfere with the function of the protein it is fused to.

The main drawback is that the affinity media is based on an immobilized antibody, and therefore has a limited binding capacity, resulting in either larger column sizes or smaller amounts purified per batch. The chromatography media also comes at a higher cost than alternative affinity media.

Key + low +++ high

Conclusion

There are many other tags that you can use but we hope this overview will give you ideas for what you should consider when choosing a tag for your protein. The key is to know your protein and the tag’s characteristics for a good choice and purification.

For more information, tips and ideas on tagged proteins, view our webinar with Professor Richard Burgess, Editor in Chief of the journal Protein Expression and Purification, and also check out this excellent collection of papers collated by Dr Richard R. Burgess which is an excellent review and includes research articles illustrating many of the purification tags in current use, including detailed experimental descriptions, example protocols, strategies and best practices for using tags.
What experiences to do you have from such tags? Have you seen other benefits or drawbacks when using the tags described here? Please use the comment field below to share your thoughts.