Immunofluorescent Localization of Tubulin

LEVEL II

Materials

 

• Coverslip cultures of an appropriate monolayer cell line
• Phosphate buffered saline (PBS)
• Acetone/Methanol (absolute) in a 50:50 volume mixture
• Rabbit anti-tubulin (or other primary antibody to tubulin)
• FITC-labeled goat anti-rabbit (or secondary antibody to match the primary)
• Fluorescence Mounting Media
• Fluorescent Microscope equipped with 490 nm excitation filter and 515 nmbarrier filter
• Kodachrome film or equivalent color slide film (Kodak Tri-X or Ilford HP400 may be substituted for black and white photography)

Procedure

 

1. Set up a coverslip culture of an appropriate cell line 24 hours prior to the lab. This is best accomplished by dry sterilization of #1 coverslips which are subsequently placed in plastic tissue culture plates. Cells are placed on the coverslips with sufficient media to cover and allowed to grow for 24 hours. There should be sufficient cells to view comfortably, but they should not be crowded on the slide.

    

2. Remove the coverslip from the culture plate and dip several times in a beaker of phosphate buffered saline (PBS) to rinse off the culture media. Drain, but do not allow to dry.

    

3. Immediately immerse in a 50:50 mixture of acetone/methanol at room temperature. Allow the coverslips to remain in the acetone/methanol for 2 minutes.

    

4. Remove the coverslips from the acetone mixture and rinse 2X with PBS.

    

5. Prepare a 1/40 anti-tubulin dilution using PBS. PbS alone may be used or better, augment the PBS with 3% (w/v) Bovine Serum Albumin (BSA).

   It may be necessary to check the appropriate antibody dilution. If so, make 1/10, 1/100, 1/1,000 and 1/10,000 dilutions to establish the correct titer. Working dilutions also may vary with the manufacturer - check the literature that accompanies your primary antibody.

    

6. Place the coverslip in a petri plate containing filter paper moistened with PBS. Make sure the cells are pointed up when placed in the petri plate! Flood the coverslip with 50 ml of 1/40 dilution of the primary antibody (or as determined in step 5).

    

7. Incubate at room temperature for 1-4 hours. The incubation may be left overnight if necessary.

    

8. Wash 3X with PBS. Place coverslips in a new petri plate containing PBS moistened filter paper.

    

9. Apply 50 ml of FITC-labeled second antibody. A 1/100 dilution usually is satisfactory. You may need to determine the appropriate dilution based on manufacturer directions or through trial and error dilutions in the range of 1/10 to 1/300.

    

10. Incubate for 30 minutes at room temperature.

     

11. Wash 3X with PBS.

     

12. Place a drop of glycerin or appropriate commercial fluorescent mounting media on a slide and place the coverslip onto the slide with the cells facing down into the glycerin.

     

13. Observe immediately with a fluorescent microscope adjusted for fluorescein (490 nm excitation and 515 barrier filter). The slides are best photographed using Kodak Ektachrome or equivalent with an ASA or 200-400. An exposure of 1-2 seconds is usually sufficient, although for low light, 30 seconds may be required. It is best to make a test exposure roll if a photometer is not available.

 

Figure 2.7 Microtubules observed via fluorescent labeling

Notes:

The procedure works for most primary antibodies merely by replacing the anti-tubulin with another appropriate antibody (anti-actin, anti-laminin, etc.). Just be sure to keep the secondary antibody appropriate to the host for the primary.