In vitro Assessment of Pathogen Effector Binding to Host Proteins by Surface Plasmon Resonance

[Abstract] The mechanisms of virulence and immunity are often governed by molecular interactions between pathogens and host proteins. The study of these interactions has major implications on understanding virulence activities, and how the host immune system recognizes the presence of pathogens to initiate an immune response. Frequently, the association between pathogen molecules and host proteins are assessed using qualitative techniques. As small differences in binding affinity can have a major biological effect, in vitro techniques that can quantitatively compare the binding between different proteins are required. However, these techniques can be manually intensive and often require large amounts of purified proteins. Here we present a simplified Surface Plasmon Resonance (SPR) protocol that allows a reproducible side-by-side quantitative comparison of the binding between different proteins, even in cases where the binding affinity cannot be confidently calculated. We used this method to assess the binding of virulence proteins (termed effectors) from the blast fungus Magnaporthe oryzae , to a domain of a host immune receptor. This approach represents a rapid and quantitative way to study how pathogen molecules bind to host proteins, requires only limited quantities of proteins, and is highly reproducible. Although this method requires the use of an SPR instrument, these can often be accessed through shared scientific services at many institutions. Thus, this technique can be implemented in any study that aims to understand host-pathogen interactions, irrespective of the expertise of the investigator.

The biochemical study of the interactions between effectors and their targets often employs qualitative techniques such as Yeast-Two-Hybrid (Y2H) (Mukhtar et al., 2011;Weßling et al., 2014) and coimmunoprecipitation (co-IP) (Fujisaki et al., 2015;Dagdas et al., 2016). However, small differences in binding between effectors and host proteins can have major impacts in function (De la Concepcion et al., 2018). Therefore, techniques that can quantitatively determine the binding between an effector and a given host protein are increasingly being required.
Isothermal Titration Calorimetry (ITC) has been commonly used as the gold standard to measure interactions between pathogen effectors and host virulence targets Maqbool et al., 2016). This technique has also been used to investigate the binding between effectors and immune receptors, and how this is translated into immune recognition (Zhang et al., 2017). However, in many cases, multiple allelic variants of both effectors and host proteins are involved in the virulence/immunity process (Zess et al., 2019), increasing the number of combinations to test and the labor-intensity of this approach. This, together with the requirement of relatively large amounts of purified proteins, can make the study of interactions by ITC impractical in some cases.
Surface Plasmon Resonance (SPR) has several advantages over ITC. First, the microfluidic nature of the technique allows the use of very small volumes of proteins at often nanomolar concentration, reducing the amount of purified protein required for the experiments compared with ITC. Also, as SPR is a high-throughput and automatable technique, multiple interactions and their respective controls can be tested at the same time under the same conditions, increasing the robustness and reproducibility of the data.
We have successfully used SPR to understand how direct binding of a domain from the rice NLR Although SPR can be used to calculate binding affinities and kinetics, this was not possible for some Pik-HMA/AVR-Pik combinations due to weak binding (De la Concepcion et al., 2018 and 2019). However, the SPR protocol presented here ranks the binding of different AVR-Pik variants to different alleles and mutants of the integrated HMA domain of the Pik receptor in the absence of precise quantification of binding affinities (expressed as equilibrium dissociation or KD values), allowing us to overcome these issues with respect to biological function (De la Concepcion et al., 2018 and 2019). Therefore, this method presents a quick way to screen and quantitatively rank interactions, which will be informative to understanding the biological implications of the interactions.
Although the protocol presented here has been optimized for the interaction between two proteins where one partner is immobilized on a Nitrilotriacetic Acid (NTA) chip through a hexa-histidine tag, it is 3 www.bio-protocol.org/e3676

Proteins of interest
The proteins to be tested must be purified and concentrated prior dilution into the running buffer.
One of the proteins whose interaction wants to be tested must contain a histidine tag while the other requires no tag.

Preparation of stock dilution of His-tagged (ligand) protein
The concentration of the protein that will be immobilized on the chip is measured and diluted in buffer (20 mM HEPES, 150 mM NaCl, pH 7.5) to obtain 2 ml of stock solution at 2 μM.

Note: The stock solution can vary depending on the experiment:
The amount of stock solution we prepare in our example is larger than needed. This is because the proteins used in this example can be produce in large amounts, are stable, and can be kept on ice for a few days.
If the protein to be tested is not produced in enough quantities and/or is not stable, we recommend preparing a fresh stock solution with a lower volume each time before running.

Preparation of working solution of ligand protein
From the stock solution, we take 25 μl and dilute it with 975 μl of running buffer to obtain a 50 nM dilution. The volume of the ligand solution will vary depending on the number of cycles that are set up, and will be indicated by the Biacore TM T200 SPR control software.
B. Manual run to estimate the amount of protein immobilized onto the chip Once the proteins that will serve as ligand are prepared, test the binding to the chip by performing a manual run. In our experiments, we aimed to use a final capture level of ligand of 250 ± 50 Response Units (RU) for each ligand to be tested. 6 www.bio-protocol.org/e3676  If multiple His-tagged protein are to be tested, the concentration to use to capture on the chip will need to be optimized for each protein.

Preparation of stock dilution of analyte protein
After measuring the concentration of the protein, dilute it in buffer (20 mM HEPES, 150 mM NaCl, pH 7.5) to obtain 2 ml of stock solution at 2 μM as described above for the protein ligand.

Preparation of working solution of analyte protein
Make serial dilutions of the protein stock to obtain final concentrations of 4 nM, 40 nM and 100 nM. The total volume depends on the number of cycles and will be indicated by the Biacore TM T200 SPR control software. Each experimental cycle consists of 4 steps described below and represented in Figure 1. This cycle can be repeated multiple times in an automated fashion using different concentrations of analyte tested over different ligands.

Chip activation
As a first step, inject a solution of 0.5 mM NiCl2 with a flow rate of 30 μl/min over Flow cell 2 to activate the chip.

Ligand immobilization
After chip activation, inject the His tagged protein (ligand) to be tested (in our case C-terminally tagged AVR-Pik effector) over the flow cell 2 (FCtest). Sixty seconds are used as the injection time to achieve a desired response at this concentration. After the ligand has been immobilized buffer is flowed over FC1 and 2 to ensure any non-specific his tagged protein is removed and a stable baseline should be achieved prior to analyte injection.

Analyte injection
Once the ligand is bound to the chip surface, inject the test analyte at a given concentration (or buffer-only control) over both flow cells (FCref and FCtest). A contact time of 120 s is used as a standard to make sure the maximum concentration is (ideally) reaching the steady state. After injection of the protein, the system switches back to buffer only flow and the bound protein will start to dissociate. Generally, 120 s of buffer only is used to see this dissociation. This region of the sensorgram can be used to evaluate differences between analyte proteins as generally tighter interactions take longer to dissociate. For each analyte, we set 3 replicates of each cycle at working concentration of 4 nM, 40 nM and 100 nM. In addition, two start up cycles were carried out using buffer only as the analyte. The total running time of 11 cycles is around 4 h for each ligand to be tested and once initiated it does not require further user intervention. Multiple runs involving different ligands and analytes can be stack together in a single run.
When the experiment is completed, the NTA chip can be removed from the instrument and stored in buffer at 4 °C until next use. The chip can be re-used multiple times for different experiments. Each time a chip is re-used the capture of nickel and his-tagged protein is checked.
As long as this is what is expected the chip can be used again. On the rare occasions a chip fails it is obvious as the nickel and his tagged protein is no longer captured.

Data analysis
To compare the results between multiple cycles, the data must be normalized by correcting the different capture levels and according to the molecular weight of the different proteins. To do this, calculate the theoretical maximal binding at saturation of the analyte (Rmax) value for each run (Buckle, 2001;Majka and Speck, 2007). This value is measured in Response Units (RU), which is how binding events are recorded in SPR. This is calculated following the equation: Once we establish the Rmax for each run, we can express the level of binding as the percentage of Rmax calculated as follows: Where RUmax is the binding response measured immediately after the end of the injection of the analyte and expressed in Response Units (RU).