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

A Swift Introduction to Electrophoresis

Electrophoresis is a very common technology used in many different types of protein analysis, no matter if the protein of interest is purified or part of a complex sample. When optimizing conditions for expression of recombinant proteins, electrophoresis can be used to obtain information about protein yield at various conditions. Additionally electrophoresis can be used subsequently to purification by gel filtration, to verify protein purity and to confirm that the purified protein has correct molecular weight.

Electrophoresis of proteins is usually carried out by loading a sample into a well, to which a voltage is then applied; the varying size, shape and charge of molecules makes them move through the matrix at different velocities. At the end of the separation, the proteins are detectable as bands at different positions in the matrix.

Gel electrophoresis, as a means to separate proteins, is usually performed under denaturing conditions imparted by the presence of the detergent, SDS, both in the sample and as a constituent of the gel and running buffer. 1.4 g of SDS will bind to each gram of protein, so that any inherent charge on the protein is masked by the coating of negatively charged detergent micelles. Denaturing gels can be run under reducing conditions, where a reducing agent such as dithiothreitol (DTT) or β-mercaptoethanol is added to the sample buffer and heated. These reagents act by cleaving disulfide bonds between cysteine residues to disrupt the quaternary and tertiary structure of the proteins, creating linear chains of polypeptides. Proteins treated in this way migrate at rates that are a linear function of the logarithm of their molecular weights.

Alternatively, denaturing gels can be run under non-reducing conditions (no sample boiling and no added reducing agent) when it is important to maintain the native structure of proteins for further analysis.

Polyacrylamide gels, both as homogenous and as gradient, are the most commonly used matrices in for separation of proteins. In a complex sample where separation is desired over a wide range of molecular weights, a gradient gel with increasing gel density should be used. In such a gel, over a given time, small proteins will reach dense regions of the gel while larger proteins will migrate within less dense regions.

Hopefully that brief of electrophoresis was helpful but for more detail and information on its use please download our free handbook guide to protein purification or ask a question via the comments section below. Thanks for reading 

Getting Started on our DHFR Project

Time to get to our DHFR project. As we have explained before our aim is to express, purify and do some characterization of DHFR while learning a few things along the way.

In our previous posts, we discussed how to plan a project, identify the key characteristics of your sample and target protein, along with reviewing the key analysis methods to be aware of. We also looked at the thinking around whether you should tag your protein or not and looked at the key/common tags.

With that in mind, a couple of things we need to consider; since we are planning to characterize DHFR using biophysical methods, we are aiming for milligram levels of active protein at a purity of more than 95 %. For our purposes, it would make sense to add a small tag, such as the Histidine tag (unlikely to interfere with our analysis methods and would simplify the purification process). However, since we are doing this to learn and demonstrate how to use protein expression, purification and analysis methodologies and the large variety of tools and techniques, we have (deep breath :) taken the decision to express and purify human DHFR without a tag. Hopefully, this will not be too complicated.

Among the methodologies we plan to use, we have already discussed CIPP – Capture, Intermediate Purification, and Polishing. Another methodology that we plan to use throughout our project is Design of Experiments (DoE). DoE is a structured approach to experimental planning that provides a framework to explore parameters that may influence the outcome of your protocol. It will help you minimize the number of experiments that you have to carry out, while maximizing the information you get out of them in order to improve or optimize your desired outcomes. As it should add logic and structure, we will try to apply DoE to protein expression, purification and any other opportunities that we may come across as our project progresses.

In our experiment, the first step is expression. To express DHFR, we have chosen the most common host, E. coli. There are many different systems that can be used (and we will review the pros and cons of the most common in a future post) but we chose E. coli as it is simple, fast, reliable, low-cost, and easy to get high expression levels (remember we need 95%). One drawback with this host is that there are no post-translation modifications, such as glycosylation. With this choice of host, there is also a chance that we will get our protein expressed in inclusion bodies. While this would require having to refold the protein, it may not be a negative thing for the purification, since the inclusion bodies precipitate and therefore can be easily isolated, plus you get very high purity of your target protein if it is expressed in IBs. 

In our next post, we will look at the gene construct, vector and the cloning of the DHFR expression vector in the E. coli host. 

Meanwhile, if you are interested in learning more about Design of Experiments for protein expression and purification, you can take a look at our handbook.