I am extracting small extracellular vesicles from cell culture media. To do that, I have harvested cell culture media from bioreactors, adherent cell cultures, and neurosphere cultures. I have only harvested the media, not the cells.
Protocol consists of harvesting medias, then
1) Spinning the collected cell culture media in a conical tube for 30 minutes at room temp at 2500g, harvest supernatant and store at 4C.
2) Spin the harvested supernatant in a 70.1 Ti Fixed-Angle Titanium Rotor in ultracentrifuge at 10k g (10400 rpm) for 30 minutes, collect the supernatant for subsequent spins and reconstitute the 10k pellet and store.
3) Spin the supernatant of 10k at 34600 rpm at 4C, for 70 minutes (110k g), discard supernatant, reconsitute pellet and
4) additional 34600 rpm t 4c for 70 minutes (110k). reconstitute the pellet and store.
The pellets were reconstituted to pbs.
The aim is to harvest a 10k and 110k small ev-containing solutions.
I did NTA analysis, which showed small EV-like results at 80-150 nm.
BUT
When I imaged my small EV:s, there is substantial background in all of my samples. All 10k and 110k samples. Here is a sample image of my EVs.
Have anyone encountered this before? What are we seeing here as the constant background? All bottels and equipment should have been sterile.
Oiva Arvola, how did you prepare samples for EM? Are they fixed/stained or not?
Susanna, I suspect that your issue is caused by PBS. Recall that 'P' is for - phosphate! Phosphate ions readily complex with salt ions that are insoluble in water at STP.
Solution? Change your diluent to a phosphate-free saline recipe!
David Ian Leavesley , it could also be a buffer issue. But what I had in mind when asking the above questions was exactly the preparation of samples for EM.
It feels like a drop of EVs' solution in PBS was dried on a grid and then applied for imaging. This is my assumption based on the EM image.
As far as I know, EVs in their native state cannot be imaged by the EM (except liquid-phase EM). Samples must be fixed before visualization using paraformaldehyde, and glutaraldehyde, followed by treatment with uranyl salts. Micrographs of fixed and contrasted samples look different.
So, the issue could be related to the EM sample preparation protocol (not (properly) fixed and contrasted sample). Additionally, sample contamination or stability issues.
Here are a few links on EV imaging:
Article Imaging extracellular vesicles: Current and emerging methods
Article Lipid-Induced Signaling Causes Release of Inflammatory Extra...
hi,
i would also like to suggest to include a labelling step here to ensure the identity of the observed structures.
I often use a protocol looking like this:
- exosome loading undiluted at RT for 15 min
- block with 0.5% BSA 10 min
- prim Ab labelling for 30 min
- wash 5xPBS 10 min total
- R&M (1:100) 30 min
- wash 5xPBS 10 min total
- Prot A Au 10nm 15 min
- wash 5xPBS 10 min total
- wash 5xH20 10 min total (to avoid uranyl phosphate precipitation)
- embedding in 0.3% Uranyl acetate in methyl cellulose
- TEM analysis
A few tips for negative stain and TEM:
1. Glow discharge carbon coated copper grids to improve adsorption by making the surface hydrophilic. The drop of pre-enriched exosomes can be as small as 5-10 µL. Cover the grids during incubation with a lid to avoid any disturbance of the grid).
2. Upconcentration of exosomes on the grids can be performed by incubating the grid on a drop of exosomes for 10 min, then remove the grid for a few min and then put it back on the same drop of exosomes again. The exosomes will stick to the water-air interface and can be “picked up” by the grid in several steps.
3. For double labeling different sizes of protein A gold are used such as 5, 10, 15 or 20 nm.
4. In order to obtain the best possible contrast and resolution several different solutions should be tested. (Aqueous Uranyl Acetate: A 1% to 3% solution of uranyl acetate dissolved in water can be used to negatively stain of many samples. The stain has a low pH so this solution is not recommended for particles that are unstable in acid conditions, Neutral Phosphotungstic Acid: A 1% to 3% solution of phosphotungstic acid is made up in water and the pH is adjusted to 7 using sodium hydroxide. This is a useful stain for many samples but is especially good for viruses that dissociate at low pH. The stain produces less contrast than the uranyl acetate. Ammonium Molybdate: Make up a 1% solution of ammonium molybdate in water. This solution has also been used to negatively stain thawed, thin cryosections of fixed cells.
5. Remove excess methyl cellulose and uranyl acetate by using a filter paper and place the grid perpendicular to the filter paper. Barely touch the grid to the filter paper and move the grid until no methyl cellulose is removed. It should only be a thin layer of methyl cellulose covering the exosomes on the grid.
Subpopulations of exosomes can be captured with Dynabeads CD9, CD63 or CD81 and be analyzed by TEM (its also possible to capture exosomes by using Dynabeads Intact Virus enrichment magnetic beads (isolation based upon charge). After isolation and washing the exosomes on the surface of Dynabeads™ can be processed using traditional TEM protocol descrived in Pedersen et al JOURNAL OF VIROLOGY,Mar. 1999, p. 2016–2026 Vol. 73, No. 3.
In brief, for conventional Epon embedding and sectioning, Dynabeads™ with exosomes can be fixed for 1 h in 1% glutaraldehyde in 200 mM cacodylate buffer (pH 7.4), washed repeatedly in aqua destillata, and incubated for 1 h in cacodylate buffer containing 1% OsO4 and 1.5% K3Fe(CN)6. Following two subsequent 30-min incubations in 1% tannic acid and 1.5% magnesium uranyl acetate, the samples were dehydrated by using ethanol and embedded in Epon. Ultrathin sections can be stained with lead citrate.
kind regards
ketil
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