Protocol for staining whole cells with PI for cell cycle analysis:

Method:

1. Harvest cells and prepare single cell suspension in buffer (e.g. PBS + 2% FBS; PBS + 0.1% BSA)
2. Wash cells twice and resuspend at 1-2 x 10⁶ cells/ml.
3. Aliquot 1 ml cells in a 15 ml polypropylene, V-bottomed tube and add 3 ml cold absolute ethanol. (The ethanol can be added forcibly by expelling from a pipette or drop wise while vortexing…determine the best method for each cell type to prevent clumping and cell loss.) 
4. Fix cells for at least 1 hour at 4^oC. (Cells may be stored in 70 % ethanol at -20 ^o C for several weeks prior to PI staining and flow cytometric analysis). 
5. Wash cells twice in PBS. (It may be necessary to centrifuge cells at a slightly higher "g" to pellet after ethanol fixation as the cells become flocculent.) 
6. Add 1 ml of propidium iodide staining solution to cell pellet and mix well. Add 50 ml of RNaseA stock solution and incubate 3 hr at 4^oC. Final Concentration 0.5ug/ml.
7. Store samples at 4^oC until analyzed by flow cytometry.

Materials:

Propidium Iodide Staining Solution:
3.8 mM sodium citrate, 50 mg/ml in PBS.
RNase A stock solution:
10 mg/ml RNase A

References:

Crissman HA, Steinkamp JA. Rapid simultaneous measurement of DNA, protein and cell volume in single cells from large mammalian cell populations. J. Cell Biol., 59:766, 1973.

Krishan A. Rapid flow cytofluorometric analysis of cell cycle by propidium iodide staining. J. Cell Biol., 66:188, 1975.

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Protocol for FASC Analysis of Cell Cycle using BrdU and PI

Labeling of Cells with BrdU:

Add BrdU (Sigma) at a final concentration of 10 mM to approximately 1 x 10⁶ cells, and incubate under the appropriate growth conditions for 15 to 60 minutes to pulse label the cells. Times may vary depending on the cell line.

Collecting the Cells:

1. Scrape or trypsinize the cells if using adherent cells. If you are interested in analyzing cell death, collect cells floating in the media as well as the adherent population.
2. Pellet the cells at 800 to 1000 rpm for 5 to 10 minutes and remove the supernatant.
3. Wash the pellet in 5 ml of 1X cold PBS and remove the supernatant.
4. Resuspend the pellet gently in 100 ml of cold PBS and keep the cells on ice.
5. Fix the cells by dropping them slowly into 5 ml of ice cold Ethanol while maintaining a Gentle vortex.
6. Place the cells at 4^oC for at least 30 minutes or at 4^oC overnight, which is the preferred method for optimal fixation.

Processing the Cells:

1. Spin down the Ethanol fixed cells and remove the supernatant leaving a small amount of liquid in the bottom of the tube (approximately 50 - 100 ml).
2. Gently vortex the cells and slowly add 1 ml of 2N HCl/Triton x-100 to denature the DNA.
3. Incubate at room temperature for 30 minutes.
4. Spin down the cells and remove the supernatant.
5. Resuspend the cells in 1 ml of 0.1 M Na₂B₄O₇, pH 8.5 to neutralize the sample.
6. Spin down the cells and remove the supernatant.
7. For each sample, make up a master mix that includes the following:
• 50 ul of 0.5% Tween 20/1% BSA/PBS
• 20 ul of anti-BrdU-FITC (Becton Dickinson)
• 5 ul of RNAse (10 mg/ml)
8. Add 75 ul of the master mix to each sample and incubate at room temperature for at least 30 minutes. Can store at 4^oC overnight, which is the preferred method for optimal staining.
9. Spin down the cells and remove the supernatant.
10. Resuspend the cells in 1 ml of PBS containing 5 mg/ml PI (Sigma) and store in the dark.
11. Analyze all samples by flow cytometry.

Solutions:

2 N HCl/0.5% Triton = 83.33 ml conc. HCl
2.5 ml of Triton X-100
bring up to 500 ml in dH₂O
0.1 M Na2B4O7 = 19.07 g sodium borate
bring up to 500 ml in dH₂O
pH to 8.5 with HCl

A paper on the advantages of amine reactive dyes for Live Dead discrimination: aminereactivelive_deaddiscrimination.pdf

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Invitrogen LIVE/DEAD cell discrimination Dyes

Dye Preparation:

The reactive dye in each kit is provided in five separate vials. Each vial provides sufficient material for staining at least 40 cell samples. However, once reconstituted, the DMSO solution of reactive dye is somewhat unstable, especially if exposed to moisture. Unused portions may be still be useful for up to 2 weeks if stored at ≤–20°C, protected from light and moisture.

1. Bring one vial of the fluorescent reactive dye (Component A) and the vial of anhydrous DMSO (Component B) to room temperature before removing the caps.
2. Add 50 μL of DMSO to the vial of reactive dye. Mix well and visually confirm that all of the dye has gone into solution.
3. Use the solution of reactive dye as soon as possible (see below), ideally within a few hours of reconstitution.

Staining the Cells:

Buffers appropriate for cell staining include phosphate-buffered saline (PBS), Hanks’ BalancedSalt Solution (HBSS), and Dulbecco’s PBS. When using an amino-reactive dye, for example the violet-fluorescent reactive dye in product L34955, Tris buffers and solutions containing sodium azide or extraneous protein should not be used for cell resuspension and washing.

1. Centrifuge a sample of tissue-culture cells in suspension. The sample should contain at least 1 × 10⁶ cells. Discard the supernatant.
2. Wash the cells once with PBS.
3. Resuspend the cells in 1.0 mL of PBS.
4. Count the cells and adjust the density with PBS to 1 × 106 cells in a 1 mL volume.
5. Add 1 μL of the reconstituted fluorescent reactive dye (from step 1.3) to 1 mL of the cell suspension and mix well.
6. Incubate at room temperature or on ice for 30 minutes.
7. Wash the cells once with 1 mL of PBS, and resuspend the cells in 900 μL of PBS.
8. Add 100 μL of 37% formaldehyde.
9. Incubate at room temperature for 15 minutes.
10. Wash once with 1 mL of PBS-BSA, and resuspend the cells in 1 mL of PBS-BSA.
11. Analyze the fixed cell suspension by flow cytometry using the appropriate excitation and detection channel (these may vary depending on instrument used):
    Violet-fluorescent reactive dye uses 405 nm excitation and ~450 nm emission (450/50 or other filter)
    Aqua-fluorescent reactive dye uses 405 nm excitation and ~525 nm emission (525/50 or other filter)
    Green-fluorescent reactive dye uses 488 nm excitation and ~530 nm emission (530/30 or other filter)
    Red-fluorescent reactive dye uses 488 nm excitation and ~ 585 nm emission (585/42, 610/20, or other filter)

Note:

If fixation is not required, then steps 7–10 above can be skipped and replaced by washing the cells twice with 1 mL of PBS-BSA, and resuspending in 1 mL of PBS-BSA.

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7-Amino-Actinomycin D ( 7-AAD) Staining of cells for Live-Dead discrimination

7-Amino-actinomycin D (7-AAD) intercalates into double-stranded nucleic acids. It is excluded by viable cells but can penetrate cell membranes of dying or dead cells.

Method:

Stain your cells as outlined in the protocol for single color or dual-color staining with FITC and/or PE-labeled monoclonal antibodies. After the last washing step resuspend your cells as usual in 1 ml of buffer for analysis. If you want to assess viability of your samples add 1-2 microliters of the 7-AAD stock solution to each tube and mix well. Keep the samples in this solution at 4^°C protected from light for approximately 20 minutes or until analysis on the flow cytometer.

Note:

This method can now be used in combination with formaldehyde fixation of samples. Samples are first stained with 7-AAD, then fixed in 1% formaldehyde that contains 2-5 microliters/ml of actinomycin D (ref. Fetterhoff et al.); see attached protocol. 7-AAD can be used for dead cell exclusion on samples that are stained with PE (phycoerythrin)-conjugated antibodies, because the emission spectra of 7-AAD and PE can be easily separated on the flow cytometer.

Materials:

A. 7-Amino-actinomycin D (e.g., Cat #129935, Calbiochem, San Diego, CA)

B. 1 X PBS with Ca2+ and Mg2+

C. Buffer: PBS (Ca2+ and Mg2+ free)
+2% newborn calf serum (or 0.2% BSA)
+0.1% sodium azide

7-AAD stock buffer:

For long-term storage, store unopened vials of 7-AAD in the freezer. Dissolve 1 mg of 7-AAD powder by adding 50 microliters of absolute methanol directly to the vial. Mix well and add 950 microliters of 1 X PBS with Ca2+ and Mg2+ to achieve a concentration of 1 mg/ml. Store solution tightly closed and protected from light at 4^°C. We have kept this solution for several months and have not observed loss in staining activity.

References:

Schmid I, Uittenbogaart CH, Krall WJ, Braun J and Giorgi JV. Dead cell discrimination with 7-amino-actinomycin D in combination with dual color immunofluorescence in single laser flow cytometry. Cytometry 13:204-208, 1992.

Fetterhoff TJ, Holland SP, and Wile, KJ. Fluorescent detection of non-viable cells in fixed cell preparations. Cytometry 14 (Suppl. 6):27, 1993.

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Protocol for the use of actinomycin D (AD) on 7-AAD stained, formaldehyde-fixed samples

Method:

Cells are first incubated with 7-AAD for approximately 20 min, spun down and washed once with 1 X PBS. Then, a 1% formaldehyde solution containing 2-5 microliters/ml of AD (F/AD) is quickly added to the cell pellet. Cells have to be stored in the cold protected from light and can be analyzed approximately 30 min after the addition of the F/AD solution. Cells are run on the flow cytometer in the F/AD solution. Samples can be stored up to 3 days without any loss in the ability to discriminate dead from live cells.

Materials:

A. Actinomycin D ( C₁) (AD, e.g., Cat# 102008, Boehringer Mannheim, IN)

B. 1 X PBS

C. Sonicator

D. Formaldehyde solution (see protocol for preparation of 2% stock solution) 

Preparation of AD stock solution (1mg/ml):

To 1mg of AD powder add:

50 microliters of ice-cold absolute ETOH, vortex

950 microliters of 1 X PBS

Sonicate the resulting solution for 10 min at 4^°C; keep the solution overnight in the refrigerator at 4^°C, protected from light before using it.

Store solution at 4^°C protected from light.

Working dilution is 2-5 micrograms/ml.