I have been trying to process neuronal cells grown as a monolayer on glass coverslips for standard TEM but my images are not to the standard I was hoping for (see attached). It looks to me like there is a general poor quality of ultra-structural preservation.
My current protocol is:
1. Aspirate media and wash cells with 0.1M cacodylate buffer.
2. Fix with 4% PFA, 1% glutaraldehyde in 0.1M cacodylate for 1 h at room temp.
3. Wash cells 3x in 0.1M cacodylate.
4. Second fix in 1% osmium tetroxide in 0.1M cacodylate for 1 h at room temp.
4. Wash cells 3x dH20.
5. Gradually dehydrate in graded series of chilled ethanol for 15 min each at room temp (30%, 50%, 70%, 90%, 100% (x3 dry ethanol)).
6. Gradually infiltrate with resin at room temp. (30% overnight, 50% all day, 70% overnight, 100% all day, 100% overnight, 100% all day).
7. Invert coverslips (cell side down) onto full Beem capsules and polymerise overnight at 55 degrees Celcius.
8. Dip sample in liquid nitrogen to remove glass coverslip.
The technique itself is very straight forward but I would appreciate any insight as to what might be causing the problems that I am encountering.
Thanks.
1. Your pictures are not loading, so not comments on their quality...
2. Do not wash specimen in cacodylate buffer before fixation (step 1)! You are killing cells (toxic buffer) before fixation. Use some other buffer.
3. Do not understand step 5: "series of chilled ethanol for 15 min each at room temp". What for chilled ethanol? Your specimen will suffer thermal shocks without any need.
Thank you for these useful tips. I have modified the file attachment so that it should open without a problem.
Hi Martin,
I agree with Vladimir's points 2. and 3. The image is loading now.
What I would suggest is also to check the culturing medium composition and then try to balance the fixation solution with regards to osmotic pressure and ionic strength of culturing medium.
It seems to me you have a lot of granularity or contamination on your section. This might result from staining process. Please, look at the following book for explanation:
Biomedical Electron Microscopy, Illustrated Methods and Interpretations, Arvid B. Maunsbach and Björn A. Afzelius ISBN: 978-0-12-480610-8, Chapter 9 - SECTION STAINING, Pages 207-228
Regards Oldrich
You are using two fixatives, PFA and glutaraldehyde, while you should use one. Try only 2% gluta in 0.1M caco and let us know ;-)!
Using combination of two fixatives is a normal procedure. I also prefer simple glut (2.5%), but many people use more complex fixatives and like them better.
Although I do not know what you want to show, to me the picture looks not bad. To improve the quality of the image, you may try the protocol I have been using for some cell types (doi:10.1016/j.ejcb.2014.08.001):
The cells were then washed and fixed with 1 ml of 2.5% buffered 0.2 M cacodylate in glutaraldehyde at 4°C overnight. Thereafter, the cells were washed five times (300 × g, 5 min at RT) in 0.5 ml of 50 mM cacodylate buffer (pH 7.2) with incubation at RT for 3 min between each washing step. The cells were again fixed with 0.2 ml of 2% buffered OsO4 containing 25% H2O, 25% 0.2 M cacodylate and 50% OsO4 at 4°C overnight. The cells were washed five times in 0.5 ml of H2O (1200 × g, RT, 5 min) with three min incubation before each washing step. The cells were further incubated with 0.1 ml of 0.5% aqueous uranyl acetate (Sigma-Aldrich) overnight at 4°C for contrast. The cells were similarly washed five times in 0.5 ml of H2O. The cells were then dehydrated by washing in 0.5 ml of 50%, 70%, 90%, 96% and 100% ethanol concentrations, respectively, with incubation for 30 min at 4°C before washing (1200 × g, 4°C, 5 min) in each ethanol concentration. The cells were again incubated in 0.5 ml of 100% ethanol at RT for 30 min and washed at 1200 × g at RT for 5 min. They were then incubated twice in 0.5 ml readymade 100% propylene oxide at RT for 30 min and washed once after the first incubation at 1200 × g, RT, for 5 min. Finally, the cells were transferred to tubes, centrifuged and embedded carefully without disturbing the cell sediment with 1:1 ratio of Epon 812 to propylene oxide overnight at RT for dehydration. After removal of the supernatant of Epon and propylene oxide on the next day, the cells were embedded again in pure Epon for 21 h at RT and transferred to 60°C for 2 days for polymerization.
Hi there, two things: 1.The osmolarity of your fix is much too high! Keep the GA at 1% but go down with the PFA - at least half. PFA does penetrate much faster than GA, but GA has better fixing properties, that is why people use mixed fixatives. 2. Your infiltration times are way too long - which resin are you using? Epoxides are reactive! For monolayers it is totally sufficient to infiltrate in the range of hours, not days. It may also help to do the Osmium postfix on ice.
Hi Irene,
Thanks for your insight. I am using low viscosity resin from agar scientific and took the infiltration times from what I use for tissue. It makes sense to use shorter times, what would you suggest? Thanks.
Hi Martin, I looked it up and for embedding of monolayers of e.g. HeLa cells we actually do not even infiltrate by using resin diluted with solvent but go right to the full resin mix (from the 100% ethanol step). You may apply the resin for 15min and then embed in fresh resin. We are using Epon, which is not even low viscosity and it works for us when we embed directly from the 100% Ethanol.
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