Sample Preparation and Imaging of Exosomes by Transmission Electron Microscopy

This protocol describes the various techniques necessary for transmission electron microscopy including negative staining, ultrathin sectioning for detailed structure, and immuno-gold labelling to determine the positions of specific proteins in exosomes.

Transcript

The overall goal of this procedure is to observe the morphology of purified extracellular vesicles and to perform specific protein localization within extracellular vesicles using electron microscopy. This method can help answer key questions in the extracellular vesicle research period, such as the categorization of the exosome and specific proteins located inside of it. The main advantage of this technique is that it can be used to observe the portion of specific proteins located both inside and outside the exosome.

The implication of this technique is to ensure diagnosis of several disease, including cancer and bacteria infections. Though this method can provide insight into the exosome, it can also be applied to other biological samples, microcellular incubation and specific protein localization. Generally, individuals new to this method will struggle because of create contamination while non-specific binding of immunogold particle.

Using standard techniques, prepare exosomes in culture. Then ultracentrifuge the culture supernatant of HCT116 cells to pellet the exosomes from the media. Once pelleted, carefully remove the culture supernatant.

To fix the purified exosome pellet, add one milliliter of 2.5%glutaraldehyde in a 0.1 molar soldium cacodylate solution at PH 7.0. Incubate the lipid bylayer vesicles for one hour at four degrees celsius. Next, remove the fixative and rinse the pellets with one milliliter of a 0.1 molar sodium cacodylate buffer solution at room temperature for 10 minutes.

Then postfix the samples with one milliliter of 2%osmium tetroxide for one hour at four degrees celsius. Remove the fixative and rinse the exosome pellet three times with the 0.1 molar sodium cacodylate buffer, changing the buffer every ten minutes. Then dehydrate the sample by shaking it for 10 minutes each in a series of graded acetone concentrations.

Next, mix up a solution of three parts acetone and one part of low viscosity embedding mixture. Replace the acetone with this mixture and incubate the sample for 30 minutes. Continue to increase the ratio of the low viscosity embedding medium, incubating for 30 minutes with each step.

Finally, incubate the exosome pellet in 100%of the low viscosity embedding mixture overnight at room temperature. The following day, embed the sample in pure low viscosity embedding mixture using an embedding mold and bake it for 24 hours at 65 degrees celsius. Using an ultramicrotome, prepare sections with a thickness of 60 nanometers.

Place the sections on a nickel grid and double stain some of them with 2%uranyl acetate for 20 minutes, followed by lead citrate for 10 minutes. Next, place the stained section into a transmission electron microscope and view the sample using 80 kilovolts and follow automatic settings for the exposure time. With an exosome image in view, acquire and save the image using the microscope software.

To begin, incubate grids containing 60 nanometer thick, unstained sections in 50 microliter drops of 0.02 molar glycine for 10 minutes to quench the free aldehyde groups. Then, rinse the sections in 100 microliters of distilled water three times for 10 minutes each. Following the last rinse, incubate the sections for one hour at room temperature in PBS containing 1%BSA.

Then, incubate the grids in 50 to 100 microliter drops of an anti-KRS antibody for one hour. Next, wash the grids five times for 10 minutes each in a drop of PBS containing 0.1%BSA. Then, transfer the grids to a drop of an appropriate secondary antibody and incubate the sections for one hour at room temperature.

Now, wash the grids five times for 10 minutes, each with a separate drop of PBS containing 0.1%BSA. Double stain the sections with 2%uranyl acetate for 20 minutes in the dark followed by reynolds lead citrate for 10 minutes. Place the stained section into a transmission electron microscope and view the sample using 80 kilovolts and image them as previously shown.

In order to perform negative staining, fix the purified exosomes following isolation with one milliliter of 2%paraformaldehyde for five minutes. Treat thin, formvar/carbon film coated, 200 mesh, copper EM grids with glow discharge for one minute, then load five to seven microliters of the exosome suspension solution on a grid and incubate it for one minute. If the concentration of exosome is too high, dilute the concentration to an appropriate level.

Immediately stain the exosomes with about 20 drops of a filtered 1%uranyl acetate solution on the surface of the EM grid. Remove the excess uranyl acetate solution from the grid by contacting the grids edge with filter paper, then quickly rinse the grid with a drop of water. Use a pair of tweezers to place the grid on the table and cover the grid partially with a culture dish.

Allow the grid to dry for 10 minutes at room temperature and then image the grid or store it in an electron microscope grid box for future observation. Start by treating the thin formvar/carbon film coated, 200 mesh copper EM grids with glow discharge for 30 seconds. Then, load five to seven microliters of the exosomes fixed in 2%paraformaldehyde solution on the grid and incubate them there for five minutes.

Next, rinse the exosomes with 100 microliters of PBS, three times each for 10 minutes. Treat the exosome containing grids with 50 microliters of a 0.5 molar glycine solution for 10 minutes to quench the free aldehyde groups. Then, transfer the grids to a drop of PBS containing 1%BSA and block for 30 minutes.

Now incubate the grids with 50 to 100 microliters of anti-PD-L1 antibody for one hour at room temperature or overnight at four degrees celsius. Wash the grid with five separate drops of PBS containing 0.1%BSA for 10 minutes each, then transfer the grid to a drop of the secondary antibody for one hour. Next, wash the grid with five separate drops of PBS containing 0.1%BSA for 10 minutes each, then wash the grid with two separate drops of distilled water.

When finished, perform negative staining with 2%uranyl acetate, as previously shown, and image the samples, or store them in an EM grid box for future observation. The exosome morphology observed here is the product of negative staining. These exosomes show a typical cup shaped morphology, which is an artifact that can occur during the drying process.

Additional information can be gained using whole mount immunogold staining, such as the location of specific proteins in the exosome. Here, the white arrows indicate the location of PD-L1. In sectioned images, the vesicles showed a lumen structure that is known as a structural feature of exosomes.

These too can be immunogold stained to identify specific proteins throughout the exosomes. Once mastered, the immunogold staining technique can be done in fibrous cells following simple preparation and sectioning if it is performed properly. Following this procedure, other method like double immuno staining can be performed to simultaneously identify the location of two different proteins.

After each development, this technique may be used in the field of extracellular vesicles to explore disease diagnoses in biopsies. After watching this video, you should have a good understanding of how to identify specific protein localization within the exosome. Don't forget they're working with fixative solutions, such as chlorhydrate, paraformaldehyde, and osmium tetroxide can be extremely hazordous and precautions such as ventilated fume hood and latex gloves should always be taken while performing this procedure.

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Journal (Biology)

Journal (Biology)

Sample Preparation and Imaging of Exosomes by Transmission Electron Microscopy

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