Fluorescence Procedures fortheActin andTubulin Cytoskeleton in Fixed Cells2

Formaldehyde Fixation

1. Fix in 4% formaldehyde (16% stock EM grade) in CBS for 20 minutes

2. Rinse in TBS

3. Permeabilize as for methanol fixation

4. Procede as for methanol fixation

Can substitute 1-2% glutaraldehyde for formaldehyde. Quench the reaction with sodium borohydride (do this 3x 1 minute, each time use freshly dissolved borohydride-just a pinch in a 1ml tube in TBS, you will get lots of bubbles). Rinse off reducing agent in TBS (3 changes in 3-5 minutes is adequate).

Staining Actin Filaments with Fluorescent-Phalloidin

1. Fix in 4% formaldehyde (16% stock EM grade) in CBS for 20 minutes

2. Rinse in TBS

3. Permeabilize in TBS-0.5% TX for 10 minutes

4. Rinse in TBS-0.1% TX (3 changes in 3-5 minutes is adequate)

5. Block in Abdil for 10 minutes

6. Incubate in fluorescent-phalloidin (1ug/ml from 1mg/ml frozen stock in DMSO) for 20 minutes in Abdil. Do not incubate for longer than 20 minutes; highly fluorescent compounds such as fluorescent-phalloidin are usually sticky and will increase background staining with longer incubations.

7. Wash in TBS-0.1% TX

8. Incubate in 1-10ug/ml DAPI or Hoesht in Abdil to stain nuclei if required for 10 minutes

9. Wash in TBS-0.1% TX

10. Rinse in TBS

11. Drain, mount, seal

12. When sealed add water to the top of the coverslip, then aspirate.

Double label experiments

In general the best fluorescence is obtained by sequentially incubating in the individual antibodies (primary, secondary, primary, secondary). It is important to titrate the concentration of antibodies or fluorescent probes. This is because if one of the stains is very weak and the other strong, any bleed through between fluorescence channels during observation makes it almost impossible to assess colocalization. (Bleed through can be minimized with the appropriate choice of bandpass excitation and emission filters. A filter that blocks the first color can also be inserted into the light path when viewing the second color). Single label controls should be initially included to confirm the general localization of test antigens. For double label experiments that include one antibody and fluorescent-phalloidin, incubate in fluorescent-phalloidin for 20 minutes in Abdil after washing off the secondary antibody.

Extraction then Fixation

Extract in CBS with 0.1% TX100 and 1ug/ml phalloidin for 30-60 seconds. Immediately add fix of choice (do not wash after extracting). Proceed as above. For a first round of experiments always compare staining to control cells that were not extracted. If you are planning a double label experiment with fluorescent-phalloidin and wish to extract before fixing- do not substitute fluorescent-phalloidin for phalloidin in the extraction for the following reasons:

• The extraction time is too short for good intensity of fluorescence.

• The extraction is so short that phalloidin does not saturate all the binding sites, so that when you incubate with fluorescent-phalloidin later in the procedure you still get good intensity of fluorescence.

• It is too expensive.

Tubulin Cytoskeleton

Glutaraldehyde Fixation: (Microtubules alone)

1. Extract cells in Microtubule Stabilizing Buffer (MTSB) + 0.5 % TX-100 for 30 seconds.

• MTSB = BRB80 + 4 mM EGTA

  COMMENT : The most troublesome aspect of this procedure is the borohydride quenching. Please try this on a test basis before wasting valuable antibody/cells! I tape a razor blade onto the frosted part of a microscope slide (dull side facing out) and this blunt edge is placed onto a porcelain coversip holder to physically block the coverslips from floating up when transferred to the quench. However, after a few tries, this is no longer a problem and the microtubules are beautifully preserved by this method. Some cellular structures may get dislodged by the borohydride although this method has been used successfully for microtubule immunofluorescence in neurons which tend to be fairly fragile. Glutaraldehyde fixation does not preserve other antigens very well and methanol appears to be the best compromise between preservation of microtubules and maintaining antigenicity of other proteins.

  Methanol Fixation: (for co-microtubule immunofluorescence)

  (NOTE: One can extract cells in MTSB + 0.5% TX-100 for 30 seconds before fixing in methanol. Extraction can often generate artifactual localizations - especially for motor proteins where it has been documented that after extraction one often sees colocalization with microtubules which is not present in straight methanol fixation. This colocalization of motors with microtubules is abolished by addition of ATP to the extraction buffer suggesting that the observed colocalization is artifactually generated by rigor binding of motors to microtubules during the extraction.)

  NOTES: General comments on double label immunofluorescence are given above in the section on actin fixation. The one problem with methanol fixation is its destruction of chromosome morphology. Methanol tends to 'puff' up mitotic chromosomes which are best preserved by formaldehyde. For centromeric antigens, we often use formaldehyde fixation and accept the poor microtubule morphology. Autoimmune sera to centromeric components, however, often require methanol fixation and then we have to accept poor chromosome morphology. As always, the conditions will need to be optimized depending on the nature of your antibody. To maximize the chances for success, for a newly generated antibody we always try methanol and formaldehyde fixation (3% formaldehyde for 15') with and without MTSB + 0.5% TX-100 extraction and compare the observed staining with all four conditions. The optimal condition for the antibody is then used for double label immunofluorescence with microtubules. Microtubule structure is poor and very variable with formaldehyde but sometimes formaldehyde ends up being necessary. Mixed formaldehyde/methanol fixative recipes have been described but we have never tried them.

1. Fix cells in -20 deg C methanol for 3'.

2. Rehydrate in TBST 3 x 5'.

3. Process for immunofluoresence as above.

4. Add glutaraldehyde to 0.5 % final. (I generally add from a 50% stock to the container with the coverslip and mix it in gently but rapidly) - Fix for 10'.

5. During fixation, make 0.1% NaBH4 (sodium borohydride) in PBS. This is used to quench unreacted glutaraldehyde which is very fluorescent if not reduced.

6. After fixation, quench for 7'. CAUTION! The borohydride will bubble vigorously and may cause coverslips to float and flip occasionally (see comment below)

7. Rinse well in PBS and process for tubulin immunofluorescence.

8. Block in AbDil for 10'.

9. Anti-tubulin for 20' - 30' (We use DM1alpha)

10. Wash 4x TBST (TBST = TBS + 0.1% Triton X-100)

11. Secondary for 20' - 30'

12. Wash 4X TBST.

13. Wash once with TBST + 1 ug/ml Hoechst. A rapid rinse will be sufficient.

14. Drain, mount and seal.