Phalloidin Staining of Yeast

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Growth and Fixation

1. Grow the appropriate yeast strain at 30˚C (or the appropriate temperature for ts strains) to a concentration of 4-8 x 106 cells/ml. This cell density corresponds to A600 =0.3-0.6, based on the general conversion of 13 x 106 cells/ml for A600 = 1. It is important not to grow the cells to high density (>2 x 107 is probably too high), otherwise cables are not present. Need 10 x 106 cells for one staining, therefore 2 ml is sufficient. Larger cultures are also convenient because the fixed cells can be stored and then stained later.

2. When cells have reached the proper density, fix by adding formaldehyde directly to the culture medium to a final concentration of 2-4 %. We use the commercial 37% liquid stock for this application. JFA showed that formaldehyde prepared fresh from paraformaldehyde does not give better results. ("Purists" insist on fresh.) Incubate 10 min at room temperature with gentle agitation.

3. Harvest the cells by spinning in tabletop at 3000 rpm (2000 gmax) x 5 min and resuspend in 1-2 volumes of 0.1 M KPi pH 7 containing 2-4% formaldehyde. Incubate 1 hr at room temperature with gentle agitation.

4. Harvest the cells and resuspend in 1-2 volumes of 0.1 M KPi pH 7 containing 10 mM ethanolamine to quench residual formaldehyde. Incubate 10 mins., room temperature. Resuspend the cells in 0.1 M KPi. Sonicate briefly (For 30 ml: setting 3, microprobe, 20 sec per DG) to disperse clumps. Spin down and resuspend in 0.1 M KPi in about 1/30th of the original culture volume. Cells can be stored at 4˚C and used for 2-3 days.

5. GFP fluorescence imaging. To image GFP and phalloidin, decrease the concentration of formaldehyde and the time of fixation. Test a series of conditions. Also, make sure that the pH of all the solutions is above 7. Formaldehyde and acidic pH appear to decrease the fluorescence of GFP. Some formaldehyde is necessary for the phalloidin staining of the actin. One needs to find the conditions that are optimal for both.


Original method used 4X quantities of cells and stain.

1. Dry down fluorochrome-conjugated phalloidin. [We have done this with a stream of gas, but want to try Speed-vac.] We use Molecular Probes phalloidin which is stored as a 3.3 µM stock in methanol (3.3µM is 300U dissolved in 3 ml methanol current bottles advise to dissolve in 1.5 ml which results in 6.6µM) . Rhodamine-phalloidin gives the best signal, but other derivatives work well and are useful for double stains in which an antibody uses rhodamine. Redissolve at 3.3 µM in PBS with 0.1% Triton X-100 (to permeabilize the cell membrane), which takes a few minutes. Need 25 µl of this solution for each sample. [Note: A. Adams (Guthrie and Fink p. 729) says 0.2-1.5 µM is OK. JFA found good staining with 1.65 µM but not with much less.]

2. Harvest ~2.5 x 106 fixed cells in the microfuge. Remove supernatant and resuspend cells in 25 µl PBS-phalloidin. Incubate 20-30 min in the dark at 4˚C with rocking.

3. Wash the cells 3 x with PBS. The washing here is actually optional because the soluble dye does not produce significant background fluorescence. If the solution dries on the slide, then this is a problem. Suspect that less washing gives higher signal from the cells because phalloidin dissociates from yeast actin relatively quickly (compared to vertebrate actin). Proceed to mount the cells and look at them under the microscope immediately. Do not let them sit overnight.

Mounting Yeast on Slides

1. Simplest. Put 2 µl of the cell suspension on a slide and add a cover slip. (Small volume pulls coverslip down tight.) Seal w/ fingernail polish. Problem with this and next method is that only a fraction of the cells are stuck on a surface and therefore holding still for long exposures.

2. Simple: Resuspend cells in mounting medium and apply to slides. Blot as much of the liquid as possible from the edges of the coverslip and allow yeast to settle before viewing.

3. More complicated but better for photography: Adhere cells to ConcanavalinA-coated coverslips. Make 0.5 cm ring on coverslip w/ rubber cement. Place thin layer ConA on coverslip, air dry, rinse w/ H2O, air dry, apply cells, wait 2-3 minutes, aspirate. Add a drop of mounting medium to a slide and mount the coverslip.Dry the edges and then use fingernail polish.

Fluorescence Imaging

1. The fluorescence of cables is weak compared to patches, at best. Longer exposures are needed to image cables. Patches may be overexposed by necessity. One may need two different images, at short and long exposure times, to properly see the cables and the patches.  

Buffers and Reagents. Want 0.01% Na Azide in Everything (except the culture medium or when using a peroxidase, such as HRP).

0.1 M KPi, pH 7. Dilute 1 M stock.

10 mM ethanolamine in 0.1 M KPi, pH 7. Add 0.6 µl of neat ethanolamine per ml of 0.1 M KPi, pH 7. The pH will rise (ethanolamine is a base) but not over 7.5.

PBS. From a 10 X stock. See recipe box.

Mounting medium: 50% glycerol in PBS, plus 1 mg/ml of either p-phenylene diamine or n-propyl gallate. The latter needs warming and vortexing to dissolve. We often have a working tube of this in the refrigerator.

ConA: 2 mg/ml in water, made 1:2000 in PhotoFlo.

Written by Jim Amatruda, 4/16/91. Modified by JAC 11/9/92 and several times later.

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