Chemistry
Using Fluorescence Anisotropy to Measure Protein-Protein Binding Affinities
Quick fact
Fluorescence anisotropy can determine a protein binding affinity (Kd) in solution without needing to immobilize either protein, making it a powerful tool in drug discovery and studying protein interactions in near-native conditions.
Why this is interesting
You know how a spinning figure skater slows down when they extend their arms? What if I told you that the same principle—rotational inertia—can tell scientists exactly how strongly two proteins stick to each other?
Read the full explanation
Understanding Using Fluorescence Anisotropy to Measure Protein-Protein Binding Affinities
Imagine shining a flashlight (polarized light) on a small, fluorescently tagged protein in solution. The molecule absorbs the light and, after a few nanoseconds, emits light of its own. Because the protein is tumbling in solution, by the time it emits, it has rotated, and its emission is depolarized—it no longer points in the same direction as the exciting light. The faster the molecule tumbles, the more depolarized the emitted light is. Now, when this small protein binds to a large protein partner, the resulting complex is much bigger and tumbles much more slowly. The fluorophore (the fluorescent tag) stays attached to the large complex, so it rotates less during its excited-state lifetime. The emitted light remains more polarized. This change in polarization—from low to high—tells you that binding occurred. By systematically adding more of the unlabeled partner and measuring how the anisotropy (a measure of polarization) increases, you generate a binding curve. The point where the anisotropy is halfway between its initial (free) and final (fully bound) values corresponds to the Kd, the concentration of the partner at which half the labeled protein is bound.
A deeper explanation
The underlying principle is that fluorescence anisotropy (r) is defined as the difference between the intensities of emitted light parallel (Iparallel) and perpendicular (Iperpendicular) to the excitation plane, divided by the total intensity: r = (Iparallel - Iperpendicular) / (Iparallel + 2Iperpendicular). This value depends on how much the molecule rotates during the excited-state lifetime (τ). The relationship is given by the Perrin equation: r = r0 / (1 + τ/θ), where r0 is the fundamental anisotropy (when no rotation occurs) and θ is the rotational correlation time, which is directly proportional to the molecular volume (ηV/RT, where η is viscosity, V is molecular volume, R is gas constant, T is temperature). When a small protein (fast tumbling, short θ) binds a large partner, the complex's molecular volume increases dramatically, lengthening θ. If the fluorophore's lifetime τ is comparable to θ (typically nanoseconds), the anisotropy will increase noticeably. The measured anisotropy is a population-weighted average of free and bound states, so a titration yields a smooth curve. The inflection point of this curve gives the Kd. Since anisotropy is a ratio of intensities, it is largely insensitive to protein concentration, a major advantage over intensity-based methods. Furthermore, this method requires only a fluorophore-labeled protein, which is usually achieved by attaching a small dye to a surface-exposed cysteine or via chemical labeling.