Size Exclusion Chromatography for Separating Extracellular Vesicles from Conditioned Cell Culture Media

1. Preparation of buffers and reagents

NOTE: Make cell culture reagents in cell culture hood to maintain sterility.

1. Preparation of cell culture reagents
   1. Prepare normal high glucose DMEM by adding 50 mL of FBS and 5 mL of penicillin-streptococcus (Pen-Strep) into 500 mL of high glucose DMEM and store at 4 °C. Use this media for culturing and expanding cells.
   2. Prepare exosome-depleted high glucose DMEM by adding 50 mL of exosome-depleted FBS and 5 mL of Pen-Strep into 500 mL of high glucose DMEM and store at 4 °C. Use this media for cell treatments prior to EV isolation.
   3. Prepare tunicamycin by adding 10 mg of tunicamycin to 1 mL of dimethyl sulfoxide (DMSO). Make 50 µL aliquots in 0.2 mL tubes and store at -20 °C.
2. Preparation of EV separation reagents
   1. Prepare 1x phosphate buffered saline (PBS) by adding 9.89 g of PBS powder to 1 L of deionized (DI) water in a 1 L graduated cylinder. Stir the solution on a stir plate until the PBS dissolves.
   2. Use the autoclave to sterilize the glassware before adding PBS. Put the glassware into a bin, close the door, and sterilize for 30 min at 121 °C.
   3. Inside of a laminar flow cabinet, connect a vacuum to the 0.22 µm filter and place filter on top of the autoclaved bottle. Slowly pour the PBS onto the filter and collect the filtered PBS in the autoclaved bottle.
   4. Prepare 0.5 M sodium hydroxide by adding 9.99 g of sodium hydroxide to 500 mL of 1x PBS in a 500 mL graduated cylinder. Stir the solution on a stir plate until the sodium hydroxide dissolves into the PBS and then filter using a 0.22 µm filter into an autoclaved 500 mL bottle.
   5. Prepare 0.05% sodium azide by adding 0.25 g of sodium azide to 500 mL of 1x PBS in a 500 mL graduated cylinder. Stir the solution on a stir plate until sodium azide dissolves into the PBS, and then filter using a 0.22 µm filter into an autoclaved 500 mL bottle.
3. Preparation of protein isolation, quantification, and identification reagents
   1. Prepare 100 mL of RIPA lysis buffer by combining 1 mL of 1 M Tris (pH 7.4), 1 mL of 10% sodium dodecyl sulfate (SDS), 1 g of deoxycholate, 1 mL of NP-40, and 0.89 g of sodium chloride (NaCl). Bring the volume to 100 mL with distilled H₂O and store at 4 °C. To make RIPA plus phosphatase and protease inhibitor solution, add 10 µL of phosphatase and protease inhibitor for every 1 mL of RIPA buffer in a 15 mL tube and store the solution at 4 °C.
   2. Immediately before use, prepare the BCA assay working reagent by adding 10 mL of reagent A and 200 µL of reagent B provided by the BCA assay kit into a 15 mL tube. Immediately before use, prepare the microBCA assay working reagent by adding 25 parts of reagent A, 24 parts of reagent B, and 1 part of reagent C into a 15 mL tube.
   3. Prepare 10x gel electrophoresis running buffer by adding 30.3 g of Tris base, 144 g of glycine, and 10 g of SDS in 1 L of DI water. Store the running buffer at 4 °C. Prepare 1x transfer buffer by adding 3.03 g of Tris base, 14.4 g of glycine, and 200 mL of methanol and adjust the volume to 1 L with DI water. Store the transfer buffer at 4 °C.
   4. Prepare Tris buffered saline with Tween-20 (TBS-Tween) by adding 2.4 g of Tris base, 8 g of NaCl, and 1 mL of Tween-20 into 1 L of DI water. Store the TBS-Tween at 4 °C.
   5. Prepare 5% blocking solution by adding 5 g of powdered non-fat milk to 100 mL of TBS-Tween. Store the blocking buffer at 4 °C.
   6. Immediately before use, prepare chemiluminescent substrate by adding 4 mL of peroxide solution and 4 mL of luminol/enhancer solution into a 15 mL tube and invert gently to mix.

2. Culturing and treating cells

1. Seed two T175 cell culture flasks with 2.3 x 10⁶ human oligodendroglioma (HOG) cells each. Bring final volume of normal high glucose DMEM to 25 mL and incubate at 37 °C with 5% CO₂ in a cell culture incubator for 48 h.
2. At the end of the 48 h, warm exosome-depleted high glucose DMEM in a 37 °C water bath. For treatment, add 25 µL of 10 mg/mL of tunicamycin to 25 mL of exosome-depleted media, final concentration of 10 µg/mL of tunicamycin to induce ER-stress. For the control, add 25 µL of DMSO to 25 mL of exosome-depleted media.
3. Remove and dispose of the normal high glucose DMEM from cells and gently rinse cells and flask with 10 mL of 1x PBS. Aspirate and discard PBS.
4. Add 25 mL of control media into the flask labeled control, and 25 mL of 10 µg/mL tunicamycin in media into the flask labeled treatment. Return flasks to cell culture incubator for 24 h.

3. Collection and concentration of conditioned CCM (Figure 1)

1. Place PBS in a 37 °C water bath. Observe flasks under a compound microscope with 10x and 100x objective lens. Take notes of any differences between the flasks. Perform the following steps for both control and treated flasks.
2. Remove media from flask and place in a 50 mL tube. Centrifuge the tube at 500 x g for 5 min at 4 °C. Transfer the supernatant to a new 50 mL tube.
3. Centrifuge the tube at 2,000 x g for 20 min at 4 °C. Transfer the supernatant to a new 50 mL tube and place on ice.
4. Obtain four 15 mL 3 kDa cutoff ultrafiltration units and add 5 mL of PBS to each tube. Prime the filter by centrifuging the ultrafiltration units at 4,000 x g for 10 min at 4 °C.
   NOTE: 3 kDa molecular weight cutoff ultrafiltration units were used in the current protocol to retain all extracellular proteins released from cells. Larger molecular weight cutoff ultrafiltration units (e.g., 30 kDa) would also work with this protocol, however extracellular proteins smaller than 30 kDa would be filtered out and removed from the samples^20,21.
5. Remove PBS from the ultrafiltration units. Divide the supernatant from each condition (control and tunicamycin treated) into two ultrafiltration units each, approximately 12.5 mL of media in each.
6. Centrifuge the tubes at 4,000 x g for 1 h 45 min at 4 °C, or until the concentrated media (referred to as the retentate) in each ultrafiltration unit is concentrated to 250 µL or less.
7. Transfer the retentate from both control ultrafiltration units into one labeled 1.5 mL microcentrifuge tube. Transfer the retentate from both treated ultrafiltration units into another labeled 1.5 mL microcentrifuge tube.
8. Measure the total volume of the retentate in each microcentrifuge tube to be 500 µL. If the sample is less than 500 µL, add 0.22 μm filtered 1x PBS until the final volume is 500 µL. Place tubes in a -80 °C freezer.

4. Cell collection

1. Place trypsin, PBS, and exosome-depleted DMEM in a 37 °C water bath. Add 7 mL of trypsin to both control and treated flasks and place in the incubator for 5 min.
2. View under the microscope to see if the cells are lifted. If the cells are not lifted, firmly tap the flask against palm of the hand to dislodge cells.
3. Wash the flask with 7 mL of exosome-depleted DMEM. Collect the cell suspension from the flask and place into 15 mL tubes.
4. Centrifuge the tubes for 10 min at 500 x g at 4 °C. Remove the supernatant and add 5 mL of 1x PBS to cell pellet. Gently pipette mix to break up the pellet.
5. Centrifuge the tubes at 500 x g for 5 min at 4 °C. Remove the supernatant and add 200 µL of RIPA plus phosphatase and protease inhibitor solution to cell pellet, pipette to mix.
6. Transfer the cells in the RIPA solution into 1.5 mL tubes. Let the tubes sit on ice for 5 min. Centrifuge the tubes at 14,000 x g for 10 min at 4 °C.
7. Transfer the supernatant to new 1.5 mL tubes. Label the tubes with treatment condition and date. Place tubes into a -80 °C freezer.
   ​NOTE: The protocol can be paused here.

5. EV separation using size exclusion chromatography (Figure 2)

NOTE: Perform the following steps twice, once for control retentate and once for tunicamycin treated retentate.
NOTE: The PBS utilized in this section is 0.22 μm filtered 1x PBS. 

1. Turn on the automatic fraction collector (AFC) and adjust settings to collect eight fractions, 0.5 mL per fraction, and buffer (void) volume to default.
   NOTE: The AFC operating conditions are as follows: column/bed volume = 10 mL; void volume = 2.70 mL; flush volume = 15 mL; optimal fraction size = 500 µL; buffer required per run = 65 mL; flow rate at 20 °C = 1.0 mL/min; reusability of the column = five uses.
2. Remove the column (see Table of Materials) from 4 °C and let the column warm up to room temperature before beginning (usually ~30 min).
3. Remove retentate samples from the -80 °C freezer and place on ice to thaw. While waiting, set eight labeled 1.5 mL tubes in the AFC carousel with lids facing inwards.
4. Insert column into the AFC with the barcode facing the reader. Start the collection by following the instructions on screen to verify there are tubes in the carousel and the waste collector is functioning properly.
5. Begin flushing the column by adding 15 mL of 1x PBS to column after the storage buffer is completely absorbed into the top portion of the column matrix. Do not add PBS too early as this will dilute the storage buffer and prevent adequate flushing.
6. Just before all of the 15 mL of PBS is absorbed by the matrix, click OK to stop the flushing and remove any residual PBS from the top of the column matrix with a micropipette. Do not touch the matrix.
7. Add 500 µL of sample to the center of the column. Click OK to begin the run. Watch the sample absorb into the matrix and quickly add 8 mL of PBS to the column immediately after the sample is fully absorbed.
8. AFC will automatically dispel the void volume into the center of the carousel and then collect all eight fractions of 500 µL each. At the conclusion of the run, remove fractions from the carousel and place on ice. Fractions 1-4 contain EVs while fractions 5-8 contain extracellular proteins. Remove void volume from the center of carousel.
9. Add 10 mL of PBS to the column after the eight fractions have been collected to wash any residual sample from the column. After the retentate sample has been fully flushed through the column, add 15 mL of 1x PBS to the column.
10. As the final PBS is being absorbed by the matrix, add 500 µL of 0.5 M sodium hydroxide to the center of the column's matrix. As the sodium hydroxide is absorbed, add 30 mL of 1x PBS to the column and let it flush through.
11. If running another sample, add eight labeled 1.5 mL microcentrifuge tubes to carousel and repeat steps 5.7-5.10.
12. Once done with all samples, complete the following steps to preserve the column for later use. After flushing 30 mL of PBS through the column as outlined in step 5.10, add 15 mL of 0.05% sodium azide to the column.
13. Leave approximately 5 mL of sodium azide on the top of the column matrix. Remove the column from the AFC and attach the top and bottom caps. Place column back at 4 °C for storage (column can be used up to five times).

6. Ultrafiltration of EV and protein fractions

1. Obtain two 2 mL, 3 kDa cutoff ultrafiltration units per sample. Use one unit to concentrate the EV fractions (fractions 1-4), and the other to concentrate the protein fractions (fractions 5-8). Each unit is comprised of three parts: cone, filter, and flow-through cylinder; label each part properly.
2. Construct the ultrafiltration unit and add 1 mL of 0.22 μm filtered 1x PBS to the filter portion and cap with the cone piece. Centrifuge the ultrafiltration unit, cone-side up at 3,500 x g for 10 min at 4 °C.
3. Remove the filtered PBS from the flow-through cylinder. Flip the ultrafiltration unit upside down (cone-side down) and centrifuge for 2 min at 1,000 x g at 4 °C. Remove any remaining PBS from the cone.
4. Add the appropriate fractions (the entire 500 µL volume) to each ultrafiltration unit (total volume will be 2 mL). Centrifuge columns cone-side up for 1 h 45 min at 3,500 x g at 4 °C, or until the retentate level is at 100 µL.
5. Remove the flow-through cylinder and discard flow through. Flip the ultrafiltration unit upside down (cone-side down) and centrifuge for 2 min at 1,000 x g at 4 °C.
6. Transfer the retentate in the cone to a labeled 1.5 mL microcentrifuge tube and bring each sample volume to 150 µL with 0.22 µm filtered 1x PBS. Place tubes in -80 °C freezer.
   ​NOTE: The protocol can be paused here.

7. Media controls

1. Obtain two T175 cell culture flasks. Label one flask control and the other treatment. Warm exosome-depleted high glucose DMEM in a 37 °C water bath.
2. Add 25 µL of tunicamycin stock solution (10 mg/mL) to 25 mL of media and put in the flask labeled treatment. Add 25 µL of DMSO to 25 mL of media and put in the flask labeled control. Keep flasks in a cell culture incubator for 24 h.
3. Collect media and process to separate EVs as outlined in steps 3.2-3.8. Subsequently, perform all steps described in sections 5 and 6.

8. Western blotting

1. Protein quantification from cell lysates and protein fractions with BCA assay
   NOTE: Protein quantification from both control and tunicamycin treated samples is performed with the following steps.
   1. Thaw lysed cells and protein fractions on ice. Make three dilution tubes for each sample; 1:2, 1:10, and 1:100.
   2. Load in triplicate on a 96-well clear-bottomed assay plate, 25 µL of bovine serum albumin (BSA) protein standards and 25 µL of each sample dilution. Add 200 µL working reagent to each well containing standard or sample, using a multichannel pipette.
   3. Incubate the plate at 37 °C for 30 min, then read plate with a plate reader at an absorbance of 562 nm. Calculate protein concentration in each sample and determine the volume of sample needed to obtain 20 µg of protein for cell lysates and 15 µg of protein for protein fractions.
2. Protein quantification from EV fractions with microBCA assay
   1. Thaw EV samples on ice. Make two dilutions for each sample: 1:50 and 1:100.
   2. Load in triplicate on a 96-well clear-bottomed assay plate, 100 µL of BSA protein standards, and 100 µL of each sample dilution. Add 100 µL of microBCA working reagent to each well containing standard or sample, using a multichannel pipette.
   3. Incubate the plate at 37 °C for 2 h, then read plate with a plate reader at an absorbance of 562 nm. Calculate protein concentration in each sample and determine the volume of sample needed to obtain 15 µg of protein for EV fractions.
3. Western blotting
   NOTE: Western blotting protocol was performed similar to as described in²², and modified as outlined below.
   1. Make 10% polyacrylamide gels with a gel making kit, using the manufacturer's instructions.
   2. For gel electrophoresis, prepare cell lysate samples by combining in a 1.5 mL tube, the volume of cell lysate necessary for 20 µg of protein, 5 µL of 4x Laemmli buffer, and the volume of RIPA buffer necessary to bring the total volume to 20 µL.
   3. Prepare EV and protein fraction samples by combining in a 1.5 mL tube the volume of sample necessary for 15 µg of protein, 5 µL of 4x Laemmli buffer, and RIPA buffer to a volume of 20 µL.
   4. Prepare media control samples by combining in a 1.5 mL tube 15 µL of sample and 5 µL of 4x Laemmli buffer.
      NOTE: Depending on the antibodies to be used, Laemmli buffer may or may not need to contain a reducing agent, such as 50 mM dithiothreitol (DTT).
   5. Boil all samples at 95 °C for 5 min, transfer samples to ice to cool, and briefly centrifuge for 30 s at 10,000 x g, then return samples to ice.
   6. Add gels to electrophoresis tank and add 1x gel electrophoresis running buffer. Load 15 µL of protein ladder and 20 µL of sample. Run the gel at 200 V for 30-50 min, or until samples have reached the bottom of the gel, just prior to running off.
   7. Transfer the protein to a PVDF membrane with 1x transfer buffer at 4 °C for 30 min at 100 V, or until the transfer is complete. Supplementary Figure 1, Supplementary Figure 2, and Supplementary Figure 3 show stain-free images of each blot after the transfer is completed.
   8. Block the membrane in 5% milk/TBS-Tween for 1 h at room temperature on a shaker. Incubate membrane in primary antibody solution overnight on a shaker at 4 °C. See Table 1 for antibody dilution information.
   9. The following day, remove the primary antibody and wash the membrane 3x with TBS-Tween for 5 min each at room temperature on a shaker.
   10. Prepare appropriate secondary antibodies in 5% milk/TBS-Tween. See Table 1 for dilution information. Add secondary antibody to membrane and incubate on a shaker at room temperature for 1 h, then wash 3x with TBS-Tween for 5 min each at room temperature on a shaker.
   11. Add chemiluminescent substrate to membrane and incubate for 5 min at room temperature on a shaker. Image blot by using colorimetric and chemiluminescent analysis. Supplementary Figure 4, Supplementary Figure 5, and Supplementary Figure 6 show uncropped images of each blot.

9. TEM imaging

1. Glow discharge²³ 400 mesh carbon/formvar coated grids to remove hydrocarbons and make the grids hydrophilic.
2. Add 5 µL of EV or protein fraction sample to grids. Incubate grids at room temperature for 3-5 min.
3. Remove excess solution by wicking with filter paper. Wash the grids with 5 µL of nanopure water and remove the excess by wicking.
4. Apply 5 µL of 1% aqueous uranyl acetate and immediately wick off. Allow grids to dry. Image grids at 120 kV on a transmission electron microscope and capture images with a charged-couple device camera.

10. Nanoparticle tracking analysis (NTA)

1. Obtain EV and protein fraction samples from -80 °C and thaw on ice.
2. Turn on the computer and the NTA instrument. Open the NTA software and it will begin to initialize automatically.
3. Select the appropriate pump that contains particle-free water and click Rinse The Instrument to rinse the cell. While rinsing, inject 10-20 mL of particle-free water into injection port on the front of the instrumentusing a syringe and avoid air bubbles.
4. Cell quality check screen will pop up; click OK and cell quality check will begin. After the check is successfully passed, the autoalignment pop-up screen will appear stating: Please fill the cell with 100 nm alignment suspension (dilution 1:250,000).
   1. Prepare dilution 1 of alignment suspension by adding 10 mL of particle-free water and 10 µL of 100 nm polystyrene beads to a tube (1:1,000 dilution). Gently invert to mix. Dilution 1 has a shelf life of 2-3 days at 4 °C.
   2. Create dilution 2 of alignment suspension by adding 20 mL of particle-free water and 80 µL of dilution 1 (1:1,000) into a tube to create the dilution of 1:250,000. Gently invert the tube to mix. Dilution 2 has a shelf life of 30-60 min at room temperature.
5. Fill cell with 2 mL of dilution 2 (1:250,000). Click OK and the program will complete autoalignment (focus) and focus optimization. The analysis tab will open up creating a profile that provides results of the cleanliness and symmetry of the measuring cell in a parabola. A pop-up screen stating: View Video Microscope is now ready for experiments, will appear; click OK and the program will return to the cell check tab.
6. Rinse the polystyrene beads from the cell by injecting particle-free water into the injection port to wash the cell. Continue washing until the number of detected particles is between 0-10 (usually between 5-10 mL). Add 1 mL of 0.22 µm filtered PBS to prime the cell for the first sample.
7. Dilute the first sample with 0.22 µm filtered PBS (start with 1:1000 dilution) and enter the dilution factor into the program. Insert 1 mL of sample into the cell and wait 5 min for particle movement to slow, as they will initially move very quickly across the screen. Set the sensitivity and shutter at 75.0 and 100, respectively.
8. The number of detected particles for ideal sample dilution is 50-200 particles. If below 50 or above 200 particles are reported, adjust the dilution of the sample. If a new dilution is required, wash the cell with particle-free water until there are 0-10 detected particles. Repeat adding diluted sample and washing until the ideal dilution is found.
9. Check all 11 camera positions to visualize that particle movement has slowed and ensure that the number of detected particles is similar between all camera positions. If the number of particles differs greatly across camera positions, add more sample.
10. Click on the Measurement tab and Run Video Acquisition. Under the video acquisition tab, enter custom name for sample and identify a safe location. Check the autosave.txt and autosave.pdf boxes to save data.
11. Set the temperature to 25 °C, click on 488 nm to set laser, and click on 11 for camera positions. Click OK to run data collection. The program will begin analyzing the number of particles and size in all 11 positions. Under the analysis tab, a bar graph will appear with diameter/nm on the x-axis and particles/mL on the y-axis.
12. After data is collected, a pop-up screen titled Positions Overview will appear providing data for each of the 11 positions. If there are more than two positions removed from analysis, repeat the procedure with more sample. If not, click on the Continue button. A PDF and txt file will open containing the results. Save the files in a safe location.
13. To run another sample, go to Cell Check tab and repeat steps 10.6 through 10.12. If done, wash cell with particle-free water until the number of detected particles is 0-10 (about 5-10 mL). Rinse cell with 1 mL of 0.22 µm filtered PBS, close the program, and turn off the computer.