Herskowitz Lab Protocol
From : RKT 7/96

Immunofluorescence for Ash1-myc


Grow cells to early to mid log phaseand determine the O.D.600. I usually harvest between O.D.600 0.5 and 2. Add 7 mL formaldehyde (37%)/50 mL cells. (If I am doing several different samples, I only grow 5-10 ml of cells per sample.) DO NOT centrifuge cells prior to fixation. Shake the cells for 5 min, set in the hood 1h with occasional swirling. Pellet cells, wash two times with dH20. Resuspend cells in an eppendorf tube. Sonicate.


For a starting amount of 10 ml cells that were at OD 0.5, resuspend the cells in 0.5 ml SP. Add 1 ul B-ME and 20 ml 1mg/ml 100T zymolyase. (I use 1 ml SP, 40 ml zymo and 2 ulB-ME if I have more than 10 ml cells at OD 0.5.) Incubate cells at 37°C for 30-45 min. Check spheroplasting under the phase microscope. Unspheroplasted cells are phase bright, spheroplasted cells are phase dark. Wash cells 1X with 1 ml SP. Resuspend in 0.5-1 mL SP. Samples can be stored at 4°C for several days (weeks?!).

Prepare slides

Do this while cells are spheroplasting. Coat each well with 15 ul of 1 mg/mL polylysine. Inc. 5 min at room temp. Remove excess polylysine. Let slides dry. Wash extensively by running distilled water over slide. Place in the 37°C incubator for ~15 min to dry.

Stick and flatten cells

Put 15 ul of the cell suspension in each well, let settle 5 min. Aspirate off supernatant using a yellow tip attached to a pasteur pipette attached to a gentle aspirator. Wash cells 2x with PBS. To do the washes, I dispense drops of the appropriate solution with a pasteur pipette onto the wells, then I aspirate the solution off as described above. Let cells dry onto the slides.

Special treatments

Methanol/acetone treatment.

Chill solutions to -20°C in groovy containers (small containers that hold up to five slides) while spheroplasting. Dip slides in -20°C methanol, 6 min. Then dip for 30s in -20°C acetone. Rinse cells 5x with PBS.


I do this and all subsequent incubations in a humidified chamber, which is a petri dish or plastic container with wet paper towels on the bottom, parafilmed closed. Add 15ml blocking solution to each well. Incubate at least 1 h at room temperature. Wash 3x with PBS to remove the detergent.

1° Antibody Incubation

Dilute antibodies in 1% BSA, 1x PBS. I use a 1:300 dilution of my batch of 9E10 antibody. For your protein of interest, you should try a range of dilutions, anywhere from 1:100 to 1:2000. (Use 1:1000 dilution of HA-11 antibody for Ash1-HA.) Spin diluted antibody for 20min in microfuge. This has to be titered to optimize signal to noise. Add 15 ml diluted antibody to each well. Be sure to include a "no 1° antibody" control. Incubate 60 min, room temp. Wash 4x with PBS.

2° Antibody Incubation

I dilute our rhodamine a-rabbit antibody 1:100. Spin diluted antibody for 20 min in microfuge. Incubate 15 ml with each well for 60 min, room temp. Wash 3x with PBS, wash 2x with dH20.

Finishing up

Add 1 drop mounting medium to each well. Gently place coverslip on slide, then press hard with your finger so the coverslip doesn't slide around. Pile paper towels and a weight on top of the slide for at least 10 min, longer (an hour or so) is fine. Seal the edges of the slide with nail polish. Let it dry. I store my slides wrapped in foil at -20°C.

Necessary Solutions

SP Buffer

1.2M sorbitol
0.1M KPO4 pH10

To make:

60 ml of 2M sorbitol
6.5 ml of 1M dibasic KPi
3.5 ml of 1M monobasic KPi
30 ml ddH20
filter sterilize


(Sigma P-1524, 1 mg/ml in dH20)

Blocking Buffer

0.1% Triton X-100
1% BSA
1x PBS

Antibody Dilution Buffer

1% BSA
1x PBS
10X PBS (use your favorite recipe)

Mounting Medium:

Fluoromount G from Southern Biotechnology Associates, Inc. Add 20 ml 1mg/ml DAPI to 1 ml Fluoromount G just before using.


Purchase from Polysciences, Inc. ,Warrington, PA 18976-2590; telephone : 1 (800) 523-2575. These are teflon coated, 25x75mm slides with 10 wells, 6mm diameter each.

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