Visualizing Intracellular Sialylation with Click Chemistry and Expansion Microscopy

We present a straightforward protocol for visualizing intracellular sialylated glycoconjugates with high spatial resolution, eliminating the need for specialized equipment. This method facilitates the study of the biosynthesis, trafficking and recycling in various pathological context. To achieve this, we combine metabolic labeling, a click chemistry, and expansion microscopy.

Super-resolution microscopy techniques like STED and STORM allow scientists to surpass the diffraction barrier, but they require expensive equipment. Recently, expansion microscope has emerged as more accessible alternative that improve resolution by physically enlarging the sample. This method can be used with any routine full resource microscopes.

It does not require super resolution microscopes, which are costly, time consuming, difficult to access and require extensive optimization. Therefore, this method is easily transferable to most laboratories. To begin, add one milliliter of medium supplemented with 500 micromolar N-azidoacetylmannosamine to the prepared cells, and incubate for 24 hours at 37 degrees Celsius under a 5%carbon dioxide atmosphere.

Then, wash the samples with one milliliter of PBS. Using 500 microliters of 5%paraformaldehyde, fix the cells on each cover slip and allow them to rest for 15 minutes at room temperature. Next, prepare a custom humid chamber using an opaque box with a lid.

Place a wet piece of blotting paper at the bottom, then lay a layer of parafilm on top. Place the cover slips on the parafilm with cells facing up. For cell permeabilization, add 200 microliters of 0.5%Triton X-100 in PBS for 15 minutes at room temperature.

Then, wash the cells three times with 200 microliters of PBS. Prepare a CuAAC reaction buffer to label the Azide-modified sialylated glycans engineered using the given components. To initiate the reaction, add 200 microliters of the CuAAC buffer to each cover slip and cover the sample thoroughly.

Then, wash the cover slips three times with 200 microliters of PBS to stop the reaction and remove excess probe and reagents. Incubate the samples for one hour at four degrees Celsius in 200 microliters of BSA blocking buffer. Add 70 microliters of the solution containing the primary antibody to the cover slips and incubate for one hour at four degrees Celsius protected from light.

Then, add 100 microliters of the fluorescent secondary antibody to each cover slip and incubate for one hour at room temperature protected from light. After washing the cover slips three times with PBS, transfer the cover slips to a six-well plate and store the samples in two milliliters of PBS at four degrees Celsius for a few days protected from light. To begin, incubate the immunostain cells with n-Acryloylsuccinimide at a concentration of 3.2 milligrams per milliliter in PBS on a mechanical shaker for one hour at room temperature.

Then, wash the samples three times with one milliliter of PBS. Now, place a 70 microliter drop of monomer solution on a piece of parafilm. Add 1.4 microliters of 10%N, N, N'N'Tetramethylethylenediamine and 10%ammonium persulfate, then mix them into the drop.

Quickly place the cover slip on the solution with the cells facing down. Allow the solution to polymerize for one hour at room temperature in a humid chamber. Next, add one milliliter of digestion buffer to the hydrogel and allow digestion to proceed for three hours at 37 degrees Celsius on a mechanical shaker.

After removing the digestion buffer, wash the gel three times with two milliliters of deionized water. Now, leave the hydrogel to expand in three milliliters of deionized water for two hours, changing the water every 30 minutes. To begin, turn on the microscope and ensure the light sources are warmed up.

Then, open the Bioimaging Acquisition Software. To create the channels, select the laser excitation wavelengths corresponding to the fluorophores and activate the lasers accordingly. Select an appropriate objective lens such as a 63x oil immersion objective with a 1.4 numerical aperture or an equivalent.

Now, place and secure a 32 millimeter diameter cover slip in a holder adapted to the microscope. Place the immunostain cell sample with cells facing down on the 32 millimeter cover slip and add a drop of deionized water to prevent the sample from drying out. Then, place the holder with the cover slip on the microscope stage above the objective.

For post expansion imaging, place hydrogel on the cover slip and perform a tile scan. Use Brightfield or Low Laser Intensity Fluorescence mode to locate an area of interest and focus it. Then, set the image acquisition parameters such as scan speed, averaging number, direction, scan mode, method, and pinhole size to achieve the desired observation.

Now, visualize the cells in the software using Live Fluorescence mode. Gradually increase the laser power and adjust the detector gain and offset to amplify the signal without introducing excessive noise, ensuring clear fluorescence visualization without over exposure. Finally, perform the image acquisition and save the data in the desired format, ensuring to include proper metadata for future reference.

Once the acquisition is complete, add a small amount of deionized water to facilitate the transfer using tweezers and transfer the sample back into the six-well plate. The average nucleus Feret diameter increased from 17.7 micrometers before expansion to 70.3 micrometers after expansion, indicating an expansion factor of four. The fibroblast marking pattern before expansion microscopy showed prominent red and green fluorescence indicating subcellular details in the Golgi apparatus, but with limited resolution, preventing in-death analysis.

After expansion microscopy, a significantly enhanced spatial resolution of subcellular structures was observed with more intricate green and red vesicles visible within the fibroblast cells.