Direct Stochastic Optical Reconstruction Microscopy of Extracellular Vesicles in Three Dimensions

This protocol employs super-resolution microscopy, specifically direct stochastic optical reconstruction microscopy, also known as dSTORM, to bypass the diffraction limit and to visualize EVs with nanometer precision in three dimensions. One notable advantage of dSTORM is its ability to directly visualize particles beneath the diffraction level of light without damaging steps that alter the biochemical nature of the EV.Many evolutionarily distinct viruses employ EV signaling, therefore, dSTORM can be employed to characterize EVs for disease biomarkers and progression, as well as to visualize individual virus particles, such as SARS-CoV2. Begin by placing affinity-purified, extracellular vesicles, or EVs, onto glass-bottom, microslide, eight-well plates in a total volume of 200 microliters, and allow them to adhere to the surface overnight at four degrees Celsius.

Without removing the existing solution from the eight-well plate, fix EVs onto the plates by adding 200 microliters of 4%paraformaldehyde in 1X PBS to the EV-containing solution in each well and allow the plates to incubate for 30 minutes at room temperature. Carefully remove paraformaldehyde and excess solution with a micropipette to not disturb the EVs. Wash the EV with 1X PBS to remove excess paraformaldehyde.

Perform the wash procedure three times. Remove excess 1X PBS. Prepare 250 microliters of dSTORM Bcubed buffer solution per sample by creating a solution of five-millimolar protocatechuic dioxygenase diluted in imaging buffer as per the manufacturer's protocol.

Add 250 microliters of the prepared buffer to each well as per the manufacturer's protocol, and incubate the plates for 20 minutes at room temperature before imaging to scavenge oxidizing molecules. The EVs can be visualized immediately or stored at four degrees Celsius for a week. To prepare the beads required for the calibration of the super-resolution microscope, dilute 100-nanometer microspheres to a concentration of 0.5%in molecular biology grade water, and pipette 200 microliters into each well of a glass-bottom, microslide, eight-well plate.

Allow the beads to settle in the wells for one hour at room temperature. Without removing the existing solution, add 200 microliters of 4%paraformaldehyde in PBS to each well to the calibration bead solution, and allow to incubate for 30 minutes at room temperature. Carefully remove the paraformaldehyde with a micropipette to not disturb the beads, and wash the beads three times with 1X PBS.

Prepare the buffer as described in the manuscript. Remove 1X PBS and add 250 microliters of the prepared buffer to each well. Allow the buffer to sit for 20 minutes before visualization.

Use the Connect the Microscope button to connect to the 3D microscope before placing anything on the stage. Add 100X oil to the objective, and place the center of the well on top of the objective. In the Acquire setting, turn on the 473-and 640-nanometer excitation lasers and click on View.

Without activating the 3D lens, view the beads under the photon saturation setting by clicking on Photon counts in the Image Display Options. Set the initial laser powers to 8.4 milliwatts for the 473-nanometer laser, and 11.6 milliwatts for the 640-nanometer laser. Decrease the focus of the laser to around minus 300 nanometer or the focal plane of the calibration beads to produce a clear resolution of the individual beads.

Once the Z-plane is focused, further adjust the laser power levels to account for variation in each field-of-view. Under the Instrument functions, complete 3D mapping calibration and channel mapping calibration to obtain the errors on the X, Y, and Z-axis. set the max number of fields-of-view to 20, the target number of points to 4, 000, the max distance between channels to 5.0 pixels, and the exclusion radius between channels to 10.0 pixels during channel mapping calibration.

Ensure that the calibration produces a point coverage of greater than 90%and mapping quality that is good. Save the given calibration data for future image acquisitions. Add 100X oil onto the objective, and place the prepared EVs onto the microscope.

Without activating the 3D lens, turn on the 640-nanometer excitation laser, and initially raise it to between 1.2 and 12.5 milliwatts, depending on the intensity of the signal in field-of-view to excite the red membrane, intercalating, dye-stained EVs. Under the Image Display Options, switch the viewing method from photon saturation to percentiles to better visualize the EVs. Adjust the laser power to minimize noise while maximizing signal and maintain all other parameters.

Adjust the focus of the Z-plane by clicking the up or down icon on the Z-axis. Set the exposure time to 20 milliseconds, the frame capture to 10, 000 frames, and the initial laser power to between 1.2 and 12.5 milliwatts, depending on the intensity of the signal and field-of-view. Activate the 3D lens using the icon, and start the acquisition by clicking on the Acquire button.

Throughout the image acquisition process, raise the laser power by three increments of 10 every 1000 frames, or enough to maintain a high signal-to-noise ratio. Do not adjust the Z-plane during acquisition. After the image acquisition, toggle over to the Analyze viewing window.

Perform drift correction on the unfiltered image, and then activate filters. Adjust photon count, localization precision, sigmas, and frame index, as mentioned in the manuscript. Overlay an X, Y, Z-plane view tool along the X-axis of individual EVs from the field-of-view and export the individual csv files of photoswitching events.

Bisect individual EVs on the X, Y-axis in an X by Y field-of-view using a Line Histogram tool, which pins photoswitching events into set distance groups. Take images of single EVs and save them as tif files. Create 3D videos of individual EVs using a 3D visualization tool and color according to placement along the Z-axis.

Following the calibration of the microscope that produced an average error of 16 nanometers on the X, Y-axis and 38 nanometers on the Z-axis, the purified u20s EVs were successfully visualized with a resolution of up to 20 nanometers on the X, Y-axis, and 50 nanometers along the Z-axis. Individual EVs visualized through dSTORM in 3D photoswitched throughout the 10, 000 frame exposure as the laser power was increased, and were readily apparent in the acquired image. Post-acquisition image correction in the Z-plane, photon counts, sigmas, and localization precision of the reconstructed image resulted in a clear resolution of the EV in 3D.

The EV photoswitched during only the first 7, 000 frames, as seen by the legend in the upper right corner. The histogram confirms that the majority of photoswitching events occurred within a 100-nanometer radius, validating that the visualized EV is an exosome, and that the isolation of EVs of a small diameter was successful. Size distribution analysis performed on other individually-traced EVs using the Line Histogram tool and X, Y, Z-plane View tool confirmed that most photoswitching events occurred within a 100-nanometer radius of the center.

The error along the Z-axis was increased, producing an elongated final image of the EV along the axial axis. Photoswitching events were not correlated with EV size, demonstrating that dSTORM-based characterization can be used for small EVs like exosomes and small, enveloped viruses less than 100 nanometers in diameter. While performing dSTORM it's important to remember to set the initial laser power very low, and slowly raise it throughout image acquisition in order to prevent photobleaching.

Since its development, dSTORM has allowed researchers to better understand the morphology of sub-cellular structures that have previously been impossible to visualize due to the diffraction limit of typical light microscopy.