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Simple yeast immunofluorescence

 

 

Application

 

This is my simplified method for doing IMF.  It’s easy and works very well for detection of tubulin Pds1-13Myc, Esp1-13Myc and Cdc14-13Myc.  I find that IMF of HA-tagged proteins is much messier, so it may require special tricks to clean up contaminating immunofluorescence.  When working with a particular protein, it may be necessary to optimize fixation time and antibody concentrations. 

 

 

Reagents and materials

 

37% formaldehyde

PBS pH 7.4

SCE pH 7.0

10mg/ml 100T zymolyase in H2O

from arthrobacter luteus

Seigaku Corporation #120493

Remove insoluble debris by spinning in a microcentrifuge at max rpm for several minutes.

multiwell printed microscope slides

            Carlson Scientific, Inc. #101205

            Polysciences, Inc., Warrington PA, #18357

concentrated HCl (12.1N)

95% ethanol

1mg/ml poly-L-lysine

            Sigma #P 8920

blocking solutions

PBS/10% NGS (normal goat serum)

PBS/1% BSA/0.05% sodium azide

primary antibodies

Serotec rat anti-tubulin tissue culture supernatant (use at 1:10 final)

9E10 mouse monoclonal anti-Myc, 0.5mg/ml in 50% glycerol (use at 1:500 final)

12CA5 mouse monoclonal anti-HA, 0.5mg/ml in 50% glycerol

secondary antibodies

rabbit anti-rat CY3, in 50% glycerol (use at 1:1000 final)

goat anti-mouse FITC, in 50% glycerol (use at 1:1000 final)

donkey anti-mouse CY3, in 50% glycerol (use at 1:1000 final)

mounting medium with DAPI

Vectashield, Vector Industries #H-1200

            Prolong, Molecular Probes

nail polish

 

 

Procedure

 

1          Collect and fix the cells

For most work, it will be necessary to collect cells in mid-logarithmic growth (~1x107/ml).  I usually collect a 1ml sample.  This is enough cells for many slides, and can even be scaled down.  Add formaldehyde to cells in growth medium to a final concentration of 3.7% and incubate the samples at room temperature to fix the cells.  Some prefer to remove the growth medium and fix the cells in water.

note: Time of fixation is a key variable in the ability to see an immunofluorescence signal. 

 

2          Wash the cells and resuspend them in SCE

Wash the cells once with water, and resuspend them in 1ml of SCE.  Sonicate the cells with the Branson Sonifier for 10 pulses, using the 30% duty cycle at output level 2.5.  Fixed cells can be stored in SCE at ~4oC for up to several weeks before processing for IMF.

 

4          Treat the cells with zymolyase to remove cell walls

Take an aliquot of cells in SCE, such as 200μl, and add an equal volume of SCE containing 10ul of zymolyase per ml  (100ug/ml stock solution).  Incubate the samples at room temperature on your bench top for 15-35 minutes.  I find 20 minutes is sufficient time to spheroplast the cells.  If cell walls are properly digested, the cells will look light gray under a microscope.  Overdigested cells will look ragged and dark gray and will not be usable for looking at cell morphology.

 

5          Wash and resuspend the cells in SCE

After the cell walls are removed, wash the samples once in SCE and resuspend them in SCE at ~¼ of the original cell volume to concentrate the cells.  Cells are fragile after cell wall removal so centrifuge them gently (1 minute at 4000rpm is sufficient), and resuspend by gentle agitation.  Do not vortex.

 

6          Prepare a multiwell slide for the samples

I use slides from Carlson Scientific.  Wash the slides in a Coplin jar with HCl (at ~1N) for 10-15 minutes, then rinse thoroughly with distilled water followed by 95% ethanol.  Allow the slides to air dry.  Put a droplet of ~10ul of 1mg/ml poly-L-lysine on each well, spread it across the well and leave at room temperature for 10-20 minutes.  Remove the excess poly-L-lysine with an aspirator, and air-dry the slide thoroughly.  Briefly rinse the slide in nanopure water and allow it to dry again.  Slides can be prepared well ahead of time and stored dry. 

 

7          Place the cells in the slide wells

Place a droplet of cells (~10ul) in each well and allow 15 minutes for the cells to adhere.  Remove any loose cells with an aspirator off to the side of the slide well.

 

8          Wash the slide wells to remove loose cells

Wash each well 2x by applying a droplet of PBS or nanopure water that is then aspirated away. Alternatively, wash by dipping the slides in PBS in a Coplin jar.

 

Option: Instead of washing the slide in PBS, immerse it in -20oC methanol for 5 minutes, followed by -20oC acetone for 30 seconds.  Allow it to dry completely.  This treatment will fix the cells to the poly-L-lysine, and may help solve adhesion problems.   

 

9          Block the samples to reduce background fluorescence

To reduce background fluorescence, block the cells by putting a droplet of blocking solution on each well, and incubating the slides for 15 minutes (room temperature is ok).  Remove the excess solution with a micropipette.  I block with PBS/10% normal goat serum (NGS), when using secondary antibodies raised in goat.  Another good blocking solution is PBS/1% BSA.  If you plan to store a PBS/BSA solution, add 0.05% sodium azide to keep it sterile.

note: Spin down diluted blocking and antibody solutions for a few minutes and use from the top to avoid aggregates.  For the remainder of the procedure, be careful not to allow the slides to dry at any point, or debris from the antibody solutions will stick to the slide.

 

10        Apply the primary antibody and incubate the samples

Dilute the primary antibody to the desired concentration in PBS/5% NGS or PBS/1% BSA.  Add a droplet to each slide well (10ul is enough to cover the well).  Place the slide in a closed container such as a petri dish with a wet kimwipe (to inhibit evaporation).  Here are some guidelines for primary antibody incubation times: 2 hours at room temperature (my standard), 1 hour at 37oC or 4oC overnight.  

note: Perform serial dilutions of antibodies to optimize the signal/noise ratio.

 

11        Rinse out the primary antibody

Remove the antibody solution with a micropipette on an aspirator and rinse the slide wells 2-3x with a droplet of PBS added to each well, using the aspirator to remove wash solution.

 

12        Apply the secondary antibody and incubate the samples

Dilute the primary antibody to the desired concentration in PBS/5% NGS.  A typical dilution guideline for IF secondary antibodies is 1:1000 final.  Add a droplet to each well and place the slide in a petri dish as in step 12.  Incubate the slide for 1 hour at room temperature.  Limit exposure of the slides to light from here on.

 

13 Rinse out the secondary antibody

Aspirate away excess antibody solution and rinse the slide wells 3x again.  Don’t let the slide wells dry out before adding mounting medium.

 

14        Add mounting medium with DAPI to the samples

Add a small droplet of mounting medium + DAPI to each well, and place a cover slip over the slide.  Gently press the cover slip down on the slide (I use a pipette to push on the cover slip over each well).  The total amount of liquid in each slide well should be enough to spread evenly over the well without coming out from under the cover slip when pressed.  If some medium emerges from the cover slip, carefully blot it away with a kimwipe, but do not spread it over the slide.

 

15 Store the slides 

Seal the coverslip with nail polish and store the slides at 4oC or –20oC protected from light to reduce fading.

 

 

 

To store antibodies: After removing an aliquot from –70oC, add an equal volume of 100% glycerol and store for use at –20oC.  This will protect the antibody from freeze/thaw cycles.  Always protect fluorophore-conjugated secondary antibodies from light. 

 

Use of antibodies: Spin down diluted antibody solutions for 10 minutes at 4oC and use off the top to avoid aggregates. 

 

Antibody dilution:

I've been using the Serotec rat anti-tubulin tissue culture supernatant primary antibody at 1:10.  Dilutions of the mouse monoclonals such as 9E10 anti-Myc and 12CA5 anti-HA will need to be optimized depending on the identity of the tagged protein, but 1:500 is a good starting point.  Our secondary antibodies such as rabbit anti-rat CY3, goat anti-mouse FITC and donkey anti-mouse CY3 work well at 1:1000 final.  For specific experiments, it may be necessary to optimize these dilution factors.

 

Some antigen-specific fixation times:

Tubulin: Punctate staining is detectable with 15-30 minute fixation, good signal at 45 minutes, improving up to 100 minutes.

Pds1-13Myc: Good signal with 15-30 minute fixation, decreasing but visible up to at least 60 minutes.

Cdc14-13Myc: Good signal with 30 minute fixation.

 

Co-detection of GFP

When detecting an antigen by IMF along with a GFP fusion, keep in mind that GFP signal generally tends to decrease with increasing fixation time.  I find my green chromosome signal from the 256LacO system is visible with up to at least 60 minutes of fixation, but is best seen with 15-30 minutes of fixation.

 

 

 

This page last modified 08/11/2011

The Stukenberg Lab and the Burke Lab are in the Department of Biochemistry and Molecular Genetics at the University of Virginia