Peptide Binding Assays: Radioligand, Fluorescence and SPR Compared
“Binds with nanomolar affinity” sounds like a single, settled fact. In practice the number depends heavily on how it was measured. A peptide binding assay can be set up to count bound ligand at equilibrium, to time how quickly a ligand arrives and leaves, or to see how well an unlabelled compound pushes a labelled one off its site. Each design has its own assumptions and blind spots. This guide compares the main formats for assay scientists and lab managers planning binding work across a series of peptides, and flags the details that decide whether two numbers can be compared.
Three questions a binding experiment can ask
Before choosing a format, it helps to be clear which question is being asked:
- How much binds, and to how many sites? Answered by saturation binding, which gives an equilibrium dissociation constant (Kd) and the receptor density (Bmax).
- How well does an unlabelled compound compete? Answered by competition binding, which gives an IC50 that can be converted to an inhibition constant (Ki).
- How fast does binding happen and reverse? Answered by kinetic methods, which give association and dissociation rate constants.
A single affinity value reported without the format and conditions has thrown away much of what is needed to judge it.
Saturation and competition designs
Saturation binding adds rising concentrations of a labelled ligand to a fixed amount of receptor until the bound signal levels off. Fitting that curve gives Kd and Bmax. Its drawback is that it needs a labelled version of the very compound being studied.
Competition binding avoids that. A labelled reference ligand is held at one concentration, and increasing amounts of the unlabelled test peptide are added to displace it. The concentration giving half displacement is the IC50. Converting IC50 to Ki, usually with the Cheng-Prusoff relationship, uses the reference ligand’s own Kd and concentration, and assumes both compounds compete for one site in a simple way. Where binding is allosteric, or more than one site is involved, that conversion no longer holds. Our explainer on Ki, IC50 and EC50 covers the distinctions.
Non-specific binding and peptide adsorption
A labelled ligand sticks not just to its receptor but to membranes, proteins, filters and plastic. Specific binding is therefore defined by subtraction: total binding minus the binding left when a large excess of unlabelled ligand has occupied the receptor.
That makes non-specific binding a working definition that shifts with the blocking agent, the filter type and the wash routine. When it makes up most of the signal, the specific component is a small difference between two large readings and carries a large uncertainty. As a rough guide, a curve where specific binding is less than half of the total deserves caution.
Peptides make this harder because many adsorb readily to surfaces, especially at the low concentrations used in binding work. Low-binding plastics and a carrier protein in buffers help, as described in our note on peptide adsorption. Losses before the peptide even reaches the well will show up as an apparently weaker affinity.
Comparing peptide binding assay formats
| Format | What it reads | Strengths | Watch-outs for peptides |
|---|---|---|---|
| Radioligand with filtration | Bound radioactivity after unbound ligand is washed away | Reference method; small label rarely alters binding | Radioactive handling; washing can strip fast-dissociating ligands |
| Scintillation proximity assay | Signal only from label held close to receptor-coated beads | No separation step; suits plate-based work | Still radioactive; bead coating adds its own background |
| Fluorescence polarisation | Slower tumbling of a fluorescent ligand once bound | Homogeneous and simple | Needs a large size difference; the dye can be large relative to a short peptide and change its binding |
| TR-FRET and BRET | Energy transfer between labelled ligand and tagged receptor | Homogeneous; can follow binding in real time for kinetics | Receptor must be tagged; label placement matters |
| Surface plasmon resonance | Mass building up on a sensor surface | Label-free; gives on and off rates directly | Immobilising either partner can block the binding face |
Kinetics, equilibrium and ligand depletion
Kd is the ratio of the dissociation rate to the association rate. Two peptides with the same Kd can have very different residence times on the receptor, one binding and leaving quickly, the other arriving slowly and staying for hours. An equilibrium method cannot tell them apart. When comparing analogs, a modification meant to tighten binding may have changed either rate, and only a kinetic measurement shows which.
Equilibrium itself is an assumption that needs checking. The time needed to reach it is governed mostly by the off-rate, so tight, slowly dissociating ligands may need long incubations. Stopping early makes affinity look weaker, and the error is largest for the best binders. The simple check is to show that the signal no longer changes with longer incubation.
A related trap is ligand depletion. The standard equations assume that binding barely lowers the free ligand concentration. If the receptor captures a sizeable share of the added ligand, often taken as more than about ten percent, the fitted constants are distorted. Reducing receptor concentration usually fixes it.
Membranes, cells and buffers
The receptor preparation changes the answer. Membranes remove the cellular context that can influence receptor conformation. Whole cells keep it but add internalisation and peptide degradation. Purified or reconstituted receptor removes both, but may not adopt its native shape.
Buffer composition matters as well. Ionic strength, pH, divalent cations and guanine nucleotides can all shift apparent affinity at G protein-coupled receptors, partly by changing the balance between G protein-coupled and uncoupled receptor states. Two labs using different buffers can obtain different constants for the same peptide on the same receptor; our article on reproducibility in peptide assays looks at this wider problem.
Running a binding screen across many analogs
Labs profiling a series of peptides often run many plates over weeks, which puts weight on consistency:
- Fix the format and conditions for the whole series, including reference ligand, incubation time, buffer and receptor preparation.
- Base stock concentrations on net peptide content, not vial weight, since counter-ions and water shift the true concentration and therefore every Ki. See net peptide content.
- Keep each compound to one lot for the duration of the screen, and log which vials fed which plates.
- Aliquot stocks in low-binding tubes so repeated handling does not change the concentration over the course of the study.
Finally, binding is only half the picture. An agonist, a neutral antagonist and an inverse agonist may show much the same affinity; a functional readout is what tells them apart. Claims of selectivity are only as broad as the receptor panel tested, a point our article on melanocortin receptor selectivity illustrates.
Peptides supplied by Bulk Peptides are for in-vitro laboratory research only, including binding and receptor studies of this kind. They are not for human or veterinary use.
