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ICC Cell Smear Protocol for Suspension Cells

This protocol is intended as a guide only, for full experimental details please read the reference provided.

1. Adjust cells to a density of 1 X 106 cells/mL, then fix by adding 4% paraformaldehyde directly to culture media for 20 minutes at room temperature.

2. Pipette 1 mL of the fixed cell suspension into a 1.5 mL microcentrifuge tube and spin down cells for 30 seconds.

3. Remove supernatant and resuspend the cell pellet in 1 mL diH2O. Spin down at top speed for 30 seconds.

4. Remove supernatant and resuspend the cell pellet in 200 μL diH2O.

5. Add 5 μL of the cell suspension to a 22 mm x 22 mm poly-L lysine coated coverslip (~3 spots per slide), then smear the suspension using the side of a pipette tip.

6. Allow all of the liquid to evaporate by placing the coverslips on a hot plate (choose low heat setting!).

7. Check for salt crystals by placing the coverslip under a microscope.  Lightly wash the coverslip with di water if crystals are present.

8.  Place the coverslips in optical grade 6-well tissue culture plates for immediate use, or store at 2 - 8 °C for up to 3 months for future experiments.


Fluorescent ICC staining protocol for cell smears


1.  Permeabilize cells as follows, depending on protein type:

a.  For detection of either nuclear or mitochondrial protein, add 1 mL TBS with 0.5% Triton X-100.  Incubate for 10 minutes at room temperature.
b.  If antibody is specific for detection of a cytoplasmic or membrane protein, add 1 mL PBS with 0.5% Tween-20 for 10 minutes at room temperature.

2. Take off permeabilization buffer and add 1 mL PBS plus 0.1% Tween-20, not letting the specimen dry out. Wash 3 times for 5 minutes before proceeding to blocking step.

3. To block nonspecific antibody binding, incubate in 10% normal serum from the host of the secondary antibody for 1 hour at room temperature. 

4. Add primary antibody (diluted in block: 10% normal serum or appropriate) and incubate overnight at 4 °C.

5. Remove primary antibody and replace with PBS. Wash 3 x 5 minutes in PBS with 0.1% Tween-20.

6. Add secondary antibody (diluted in block: 10% normal serum) at appropriate dilution as per manufacturer’s recommendations.  Incubate for 1 hour at room temperature

7. Remove antibody and replace with PBS, wash 1 x 5 minutes in PBS with 0.1% Tween-20.  For second wash, add Hoechst 33258 to PBS at 1:25,000 and incubate for 10 minutes.  Wash a third time, adding phalloidin at 1:40 diluted in PBS with 0.1% Tween Tween-20 for 20 minutes (total of 3 X 5 minute PBS Tween-20 washes).

8. Carefully remove the coverslips from the wells and blot to remove any excess water. Dispense 1 drop of anti-fade mounting medium onto the microscope slide per coverslip. Mount the coverslip with the cells facing towards the microscope slide and tack edges down using clear nail polish.

9. Visualize using a fluorescence microscope and filter sets appropriate for the label used (please see list below). Slides can also be stored in a slide box at < -20 °C for later examination.

Fluorescent Dyes λabs (nm) λfl (nm)
Atto390 390 479
Atto425 436 484
Atto465 453 508
Atto488 501 523
Atto532 532 553
Atto565 563 592
Atto590 600 627
Atto594 601 627
Atto633 629 657
Atto637 630 659
Atto655 663 684
Atto680 670 710
Atto700 700 725
DyLight 488    493 518
FluoProbes®547H 557 572
FluoProbes®647H 653 675
FluoProbes®682 690 709
FluoProbes®752 748 772
AMCA   346 442
FITC 485 535
Rhodamine 544 576
Texas Red 595 615