There's a little trick I found in the literature that works surprisingly
well, although I'm not positive it will be exactly what you're looking
Wash some yeast cells with water, resuspend in water and pH to 4.0
(I used HNO3 for this; I doubt it matters). Mix an equal volume of the pH
4 yeast suspension with an equal volume of 0.74 mM Al(NO3)3 (aluminum
nitrate) and rotate at RT 1 hr.
Either "spot" a small volume of the aluminum-treated yeast on the
coverslip, or submerge the coverslip in the suspension;
Allow the cells to sit for 20 min at RT.
Wash with water to remove unbound cells.
This works well and is really easy. The cells are alive and I've watched
them bud several times. They may "migrate" around a little on the
coverslip, but you can actively flow water over them and they'll stay
attached. You can even use buffers of varying pH and they'll stay
attached, but only AFTER immobilization -- the low pH during the
attachment is important. I got my Al(NO3)3 from Aldrich.
This is all based on the following reference, and the chemistry behind it
is best summarized in this quote from that paper:
"Adsorption of aluminum ions decreases the negative electrostatic
potential near the yeast surface and allows adhesion of the cells to a
negatively charged support such as glass."
van Haecht, et al. Biotechnology and Bioengineering 27: 217-224 (1985).
On 25 Aug 2000, Stefan Jakobs wrote:
>> we are trying to attach living S. cerevisiae on coverslips for
> microscopy. Any ideas (other than poly-L-lsine and agarose?
> Any suggestions are highly welcome!
>> Thanks, Stefan.
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