Immunofluorescence for Ash1-myc
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.
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?!).
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
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
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.
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.
60 ml of 2M sorbitol
Polylysine(Sigma P-1524, 1 mg/ml in dH20)
0.1% Triton X-100
Antibody Dilution Buffer
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.