Fluorescence Fluctuation Spectroscopy to Study Protein Homo-Oligomerization

Fluorescence fluctuation spectroscopy can determine the oligomerization state of proteins in a sample.

To begin, take a chambered slide containing a fluorescently-labeled protein monomer solution. Place the slide under a confocal microscope. Focus the laser beam on a small part of the sample — the confocal volume — to excite the protein monomers.

The protein molecules diffuse in and out of the confocal volume due to Brownian motion. This movement causes rapid changes in the fluorescence intensity, leading to fluctuations in the fluorescence intensity over time.

Add the dimerizing agent — a bivalent ligand that helps two protein monomers bind and, thus, form a dimer. Due to dimerization, two fluorescent labels come together to form a single particle, which increases the fluorescence intensity per particle — the molecular brightness.

The increase in molecular brightness due to dimerization increases the amplitude of the fluorescence fluctuations — as two fluorescent labels enter and leave the confocal volume together.

Obtain images of the confocal volume across time before and after adding the dimerizing agent.

Compute the brightness of the individual pixels in the confocal images, and obtain the mean brightness curve over time.

The addition of the dimerizing agent results in a two-fold increase in the brightness due to the dual fluorescence signals from each particle, while the total number of protein molecules remains the same — indicating the formation of dimers.

To set up the multiwell plate array, first, prepare a solution of 100 nanomolar purified FKBP12 in the same buffer used for size exclusion chromatography. Sonicate and centrifuge with a quick spin of 13,000 RPM to prevent the formation of aggregates.

Now, pipette 100 to 200 microliters of the diluted protein into an 8-well observation chamber with a glass bottom. Add the BB dimerizer to final concentrations of 10, 20, 40, 80, 100, 150, 300, and 500 nanomolar. As a reference, prepare solution of 100 nanomolar mVenus alone to evaluate potential aggregation and precipitation effects, and to recover a brightness value for the monomer with the same acquisition settings.

Any light scanning microscope confocal system equipped with digital detectors or well-characterized analog detectors, and capable of keeping a constant dwell time for every pixel acquired can be used. Select the collar correction water immersion objective designed for fluorescence correlation spectroscopy.

Now add a drop of water to the collar correction water immersion objective. Mount the 8-well observation chamber into the stage. Set the excitation beam path by turning on the 514-nanometer laser, and setting it at 20 to 100 nanowatts power at the exit of the objective. Turn on one HyD detector. Detectors capable of photon-counting are preferable. Select the emission window from 520 to 560 nanometers.

For the acquisition mode, use 16 by 16 pixels.

Set the pixel dwell time such that the frame time is longer than the protein diffusion, and the pixel dwell time is much shorter. This corresponded to setting the dwell time to approximately 13 microseconds for the system used in this demonstration.

Set the pinhole at one Airy unit for the corresponding emission of approximately 545 nanometers. Select the xy-time acquisition mode and select the number of frames to be acquired per acquisition and well. Now, set the pixel size at approximately 120 nanometers.

If the system is equipped with high-throughput mode, introduce the coordinates of each well, and the number of acquisitions per well to automate the process. If the system is equipped with a perfusion system, load the BB solution, and program the perfusion to start right after the 5,000th frame to evaluate the kinetics of dimerization while acquiring 10,000 images.

Select the correct well, and focus on the solution. Then, start the acquisition and save the resulting stack of images in TIFF format.