Resins & Protein Binding Capacities

Compare & Contrast 

When choosing what resin to use for purifying a protein, one of the most important characteristics to look at is protein binding capacity.  Protein binding capacity is a critical parameter for chromatography media because it determines how much media is needed in order to purify a certain amount of protein. 

This in term determines the column size needed, flow-rates of your chromatography system, and ultimately the total costs for purifying your protein. But, can capacity data reported from different vendors always be directly compared?

The short answer is "No" and in this post we will try to explain what protein binding capacity is, how it can be measured, and what to look out for when looking at your vendor’s specification sheets.

Firstly, the resin capacity for different proteins is often specified by suppliers based on

• different modes of measurements (dynamic or static)

• experimental conditions (pH, salt/conductivity, protein concentration)

• reference (capacity per milliliter wet resin or g dry resin)

Naturally, what protein has been used to determine the binding capacity is vital information in order to be able to compare specs. Unfortunately, in many cases, the method for determination of the binding capacity is not stated.

This makes it very difficult to compare the resins based on tabular values from the vendors.

What are the differences between static and dynamic capacity?

The static binding capacity (SBC, also called total protein capacity) is normally measured in batch mode in a beaker and is usually referred to as the maximum amount of protein bound to a chromatography medium at given solvent and protein concentration conditions. The size of SBC   varies significantly depending on the protein loaded. In these experiments, an excess of protein is loaded to give a maximum binding capacity. Protein loss is often over 50 %.

Dynamic binding capacity (DBC), on the other hand, is the binding capacity under operating conditions (i.e. in the packed affinity chromatography column during the sample application and washing procedures). The DBC of a chromatography medium is the amount of target protein that binds to the medium  under given flow conditions before a significant breakthrough of unbound protein occurs.

DBC is determined by loading a sample containing a known concentration of the target protein. The load of the protein sample on the column is monitored and will bind to the medium to a certain break point before unbound protein will flow through the column. From the breakthrough curve (see fig.1) at a loss of for example 10 % protein (named QB₁₀),  the DBC is found and the experiment is stopped. As this parameter reflects the impact of mass transfer limitations that may occur as flow rate is increased, it is much more useful in predicting real process performance than a simple determination of saturated or static capacity.

In most instruction manuals from GE Healthcare, you can find information on the DBC and how it has been determined.

How does GE determine its resin binding capacity?

GE Healthcare uses the DBC as the measurement for media packed in columns since it takes the flow rate and bed height of the column into consideration. It also takes into account ligand density, size of protein, and the porosity of the media which are the only factors included in the measurement of SBC.

SBC can be used to determine the binding/total capacity of bulk media.

However, as DBC is measured under operating conditions you will also get information on what the maximum load of your target protein to your column should be in order to avoid unnecessary loss.

If you have any questions please ask away via the comments section or for more detail check out our Protein Skills Handbook which covers this and many other aspects of protein science

When should I use a tag to purify my protein? (and when not?)

Sometimes we’re not particularly interested in purifying proteins. We just want to get it over and done with so as to get on with the experiments that will help us understand it role. This is where tagging your protein with something that adds biospecific affinity comes in handy. It allows you to simplify the purification protocol greatly, sometimes to the extent that you can use a standard protocol.

Remember though, tagging may not always be the right solution as adding a tag can introduce changes compared to the native protein, leading to undesirable effects. For example, if you are interested in drawing conclusions about function, having a tag may introduce uncertainty, or worse,  totally destroy or alters the function of your target protein. 

Also the tag itself can interfere with your ability to use the protein the way you want; important in using therapeutic proteins, where you want to be as close to the native form as possible. In most situations though tags, especially small ones like a His-tag, have no negative affects in terms of biophysical characterisation.

If you need a protein in native form

If you have decided that you need to have your protein in the native form, you can express it without a tag and go through the process of designing a protocol involving multiple chromatographic purification steps. It can be a little tedious and potentially unnecessary. Nowadays, with so many proteins already having been expressed, it is likely that you will be able to find existing scientific literature with information on your purified your protein, or an analogue.

As an alternative to expressing and purifying the native form, you can tag your protein and then remove it later. Whilst this simplifies the purification protocol, you’ll have to figure out what tag to use and how to remove it. You will also have to add the steps for cleaving the tag (manually removing it) to your protocol. It does not always have to be manual; there are systems such as AKTA pure that can automate this for you.

Removing the tag sometimes  isn’t very straightforward. It always includes using a sequence specific protease. The blood factors Thrombin & Factor Xa are most commonly used for this, but you need to make sure that their respective cleavage sequences (LVPR↓GS and IEGR↓) are not present in your target protein. Some tags, such as the Glutathione-S-Transferase (GST)-tag can be removed with a more specific proteases, such as PreScission protease (human rhinovirus 3C protease, cleavage sequence LEVLFQ↓GP), and this is extremely unlikely to effect your protein.

If you don’t need a protein in its native form

In addition to using a tag to simplify the purification process, there are actually a number of other benefits

• A tag can add the ability to use a generic detection method (such as a standard tag-specific western-blotting antibody, or an enzymatic assay in the case of the GST-tag)
• It may actually help overcome some challenges with your protein e.g. by stabilizing it or making it more soluble, as shown in this (very cool :) example of how a challenging spider silk protein was purified by using a special solubility tag in addition to a Histidine-tag
• Some tags can be fused to proteins for a broad range of other applications such as—labeling for imaging and localization studies, protein–protein interaction studies, and subcellular localization or transduction
• Tags also allow strong binding to chromatographic affinity media in the presence of denaturants which makes it possible to purify a protein that requires this, e.g. if it has been expressed in inclusion bodies. In this case, you can then also try to perform refolding of the protein while it is still bound to the column (too complex to describe here, we will discuss this in-depth in a future post..)

Pros	Cons
Using a tag
Simple, generic purification using affinity chromatography (AC) as a first step	Tag may interfere with protein structure and affect folding and biological activity
Tags are easy to detect, in comparison to the target protein, which allows for a generic detection method	If tag needs to be removed, cleavage may not always be achieved at 100 % and sometimes some amino acids may be left
Solubility and stability can be improved	Only some tags can be used under denaturing conditions
Purifying a native protein
Tag removal is not necessary	The purification and detection protocols need to be designed specifically
Purification can always be done under denaturing conditions	Problems with solubility and stability may be   difficult to overcome and may need special protocols to be developed

       

In summary, tag if you can – it simplifies your purification work massively, but don’t be afraid to express and purify your protein in its native form. 

Let us know your experiences working with tags, preferences and if you have any questions in the comments. Next time, we will look at the pros and cons of the most common tags used for protein purification.