Frequently Asked Questions

We do NOT collect User Data during CMI training sessions.

All Training sessions include:

• Overview of the instrument and maintenance procedures
• Tutorial on control and analysis software
• Review of experimental design
• User trainings may be individual or small group, depending on scheduling.

For some technologies, we'll run a standard during training to illustrate how the instrument works and/or to practice. We won't collect any data during training on systems for which sample manipulation is not needed for effective training and for which consumables are expensive or data collection is very time consuming.

For a basic nanobody selection campaign, 1 ml antigen at 15 μM is needed. We require SEC-purified protein (and can purify your antigen by SEC if you cannot). See CMI Service Protein Requirements.

The CMI has four primary technologies to quantify molecular interactions: Surface Plasmon Resonance (SPR) and Biolayer Interferometry (BLI), Isothermal Titration Calorimetry (ITC) and MicroScale Thermophoresis (MST). Each technologies has its advantages and disadvantages. 

Some factors to consider when choosing a binding method include:

• need for kinetic and/or equilibrium fits
• the relative size of the interactants
• the amount and concentration of samples
• the ease of labeling and/or immobilizing interactants
• the affinity of the interactants

There is no one best method for measuring molecular interactions. Request a consultation at the CMI to discuss which method might be best for your specific project.

For Nanobody selections, the CMI currently uses a synthetic yeast display library, developed in the Kruse lab, a based on a consensus framework derived from llama antibody genes with variable complementary determining region loops (CDRs) designed from known nanobody structures.¹

This library is available for non-profit research from Kerafast: EF0014-FP. For-profit organizations can license the library from the Harvard Office of Technology Development.

1. McMahon, C. et al.Yeast surface display platform for rapid discovery of conformationally selective nanobodies. Nat Struct Mol Biol25,289–296 (2018).

The choice of which molecule to keep constant and which to vary will depend on the experiment type, the properties of the molecules and the stoichiometry of the interaction.

For ITC, where neither protein is labeled or immobilized, either molecule can be in the cell or syringe. You will need a higher concentration and total amount of the molecule in the syringe, so solubility and availability will generally be deciding factors. For protein/small molecule interactions, it is most common to put the protein in the cell and the compound in the syringe.

For MST, one of the molecules must be fluorescently labeled. You'll first start by determining which molecules can be easily labeled with a fluorophore.  If both molecules can be labeled (as with protein/protein interactions), your choice will generally be based on solubility and availability. You will use a constant low concentration (typically nM) of the fluorescent molecule and much higher concentration of the non-fluorescent binding partner (100X Kd).  For protein/small molecule interactions, it is most common to fluorescently label the protein and have a variable concentration of non-fluorescent compound.

For SPR and BLI, one molecule must be tethered to the biolayer surface. There are variety of ways to capture proteins onto SPR and BLI sensors, including covalent capture and many affinity capture options.  For protein/protein interactions you could immobilize either molecule.  However, it's imporant to consider stoichiometry of binding. If one of the molecules is multivalent (an antibody, for example), then you should always try to immobilize the multivalent molecule, as the fitting is much more straightforward. Other considerations are solubility, availability and existing tags for immobilization. For protein/small molecule interactions, it is most common to immobilize the protein and use a variable concentration of compound.

The amount (volume and concentration) of sample that you need will vary by method and by the Kd (equilibrium dissociation constant) of the interaction.  For guidelines see the CMI technology comparison table.  Binding assays will typically have one binding partner at a fixed concentration (or immobilized on a surface) and one binding partner at a variable concentration. The high concentration for the variable analyte will typically be 10-100X Kd, depending on the method.

The more pure your sample is, the easier it is to interpret the results, as contaminating proteins may contribute to your overall signal in bulk measurements.

Differential Scanning Fluorimetry (DLS), Circular Dichroism (CD) and Dynamic Light Scattering (DLS) signals are concentration dependent and additive. So generally, minor contaminants will be expected to contribute a low signal. However, there might be specific cases where the contaminate contributes more than you might otherwise expect to the overall signal, due to protein structure or mass. When comparing samples, similar purity and concentration between the samples will help mitigate potential artifacts.

The amount of protein needed will vary by method and sometimes by the mass of protein to be characterized. See CMI Service Protein Requirements for estimates and recommendations for the amount of protein needed for standard data collection.

If your membrane protein has fluorescent residues (tryptophan and tyrosine), you should consider trying our Prometheus NT.plex for DSF using intrinsic protein fluorescence (NanoDSF).

However, if you wish to do conventional DSF (dye-based), please note that many hydrophobic dyes, such as Sypro Orange, cannot be used for Differential Scanning Fluorimetry of membrane proteins, dues to high background staining. However, if your membrane protein has buried cysteines, then a cysteine-reactive dye may work. BODIPY FL-cysteine can be used, since it has excitation and emission wavelengths within the standard range of many qPCR instruments, including our QuantStudio 6/7.  CPM (a courmarin derivative) was one of the earlier dyes used for this purpose, however many modern qPCR instruments no longer have excitation filters appropriate for this dye.

General Experimental Conditions for Conventional DSF using BODIPY FL-cysteine:

• Alkylate exposed free thiols with Iodoacetamide prior to adding cysteine-reactive dye, so as to block background staining (1mg/ml iodoacetamide).
• Label with BODIPY FL-cysteine at a final concentration of 2 µM dye.
• 25 °C - 99 °C temperature ramp, with ramp rate of 1 °C/minute.
• 470 +/- 15 nm excitation and 520 +/- 15 nm emission filters. 

References:

• Bergsdorf, C. et al. An Alternative Thiol-Reactive Dye to Analyze Ligand Interactions with the Chemokine Receptor CXCR2 Using a New Thermal Shift Assay Format. Slas Discov 21, 243–251 (2016).
• Wingler, L. M. et al. Angiotensin and biased analogs induce structurally distinct active conformations within a GPCR. Science 367, 888–892 (2020). Adaptation by CMI User, Meredith Skiba from the Kruse lab. 

The relative size of the molecules isn't generally the deciding factor. Large proteins will typically give larger signals, but that isn’t necessarily better.

For protein/protein interactions, start by considering other properties of the proteins. Most importantly stoichiometry of binding. If one of the molecules is multivalent (an antibody, for example), then you should always try to immobilize it, as the fitting (to a 1:1 model) is much more straightforward. If both molecules are multivalent, try to immobilize at a very low density. If necessary you can fit to a bivalent analyte model. Other considerations are solubility and existing tags for immobilization.

For protein/small molecule interactions, you will typically immobilize the protein, as immobilization of small molecules is often difficult and or disruptive to binding. For small molecule interactions, a very stable capture method (e.g. biotin-capture) is recommended.

MST experiments on the Monolith NT.115 pico require that one of the binding partners is fluorescently labeled. 

For protein/protein interactions, start by considering the amounts, solubility and sequence of the proteins. The labeled target is typically used at 5-10 nM concentrations (200 ul/titration).  The unlabeled ligand is ideally used at a concentration of 100X Kd (equilibrium dissociation constant), so could be uM to mM for weaker binders (20 ul/titration).  If one protein is much more soluble and/or more abundant, that may be a better choice for the unlabeled ligand.  If one protein has a convenient labeling tag (such as His-tag or sortase tag).

For protein/small molecule interactions, you will typically label the protein, as labeling of small molecules is often difficult and or disruptive to binding. 

Unlike fluorescence polarization, the relative size of the molecules isn't generally the deciding factor when choosing which molecule to label for an MST experiment.