Houston Methodist. Leading Medicine.
Houston Methodist. Leading Medicine

Methods

Flow Cytometry Methods


Cell Cycle Analysis

Cell cycle analysis can be performed with viable or non viable cells. However, cell cycle analysis of viable cells with reagents such as Hoechst 33324 (which requires UV excitation) has been somewhat disappointing unless one is using a quickly cycling cells.   

The easiest cell cycle analysis protocol involves ethanol fixation, staining with PI (Propidium Iodide), and digestion with RNAse-A.   A copy of the protocol from Current Protocols in Cytometry is available for download here: DNA Cell Cycle Protocol.


Cell Labeling

The rule of thumb is to use 2-3 μg of primary antibody per 2-300,000 cells in a total volume of 100 μl, and to keep these ratios consistent between samples and experiments. Protocol details are as follows.

METHOD#1

1) Resuspend 3-500,000 cells in a final volume of 100 μl of 1% BSA-PBS
2) Add 2-3 ug per 1-200,000 cells and incubate on ice for 45 minutes.
3) Wash the cells three times with 1% BSA-PBS.
4) Resuspend cells in 100 μl of 1% BSA-PBS, add 2-3 ug of FITC labeled second antibody and incubate 45 minutes.
5) Wash three times with PBS without BSA.
6) Fix with 1% paraformaldehyde for 2 minutes.
7) Finally, wash one more time with PBS and resuspend in ~300 μl of PBS.

This standard protocol has low background, but cells will invariably be lost cells through all the washing steps. To minimize cell loss, use 15 ml polypropylene tubes because cells will stick to the walls of polystyrene tubes, even after centrifugation. Due to cell loss during the washing steps, we recommend starting with at least 500,000 cells in order to yield 200,000 for analysis on the flow cytometer.

METHOD#2:

The standard protocol has ben improved to reduce hands-on time and reduce cell loss during sample processing. In this version, which is now standard in our core facility, the individual washes are replaced with one large volume wash. This protocol  results in minimal cell number loss:

For the FACS Calibur/LSRII, and Fortessa, use 12 X7 5 polypropalene snap-top tubes (Fischer #14-9591-0AA - Still available from Fisher as of 09/15/2011). With the tubes labeled, the tops are removed and retained. They are replaced after step 6.   Due to differences in the molding process, not all 12 X 75 tubes are exactly alike and only certain ones work with the FACS Calibur or Vantage.   Becton-Dickenson recommends a polystyrene 12 X 75 tube that is only sold through Falcon, and works well on both machines. However, polystyrene is fairly brittle and if cracked at the lip, it will not work.  1-2% BSA-PBS can suffice for complete HEPES buffered tissue culture media in this protocol.  Label your cells in whatever media keeps them happy.   DO NOT do everying in just PBS or you will have very few cells alive before heading to the cytometer.

1) Resuspend 3-500,000 cells in a final of 100 μl of complete media.
2) Add 1-2 μg of primary antibody and incubate for 30 minutes on ice.
3) Wash once, adding 4 ml of complete media, and centrifuge 500 X g for 5-7 minutes.
4) Add 2-3 ug of FITC labeled secondary antibody in 50-100 μl of complete media and incubate 30 minutes on ice.
5a) Wash once, adding 4 mls of complete media, centrifuge 500 X g for 5-7 minutes.
6a) Resuspend in 300 μl of complete media for live analysis - keep on ice and analyze within 1-2 hours.

If fixation is needed:
5b) Wash with 4 ml of PBS (no BSA), and centrifuge 500 X g for 5-7 minutes.
6b) Resuspend in 300 μl of 1% paraformaldhye (Can store in the dark and at 4°C for 3-4 days).

Note: If the primary antibody is a hybridoma culture supernatant, simply resuspend the cells in 50-100 μl of culture supernatant, rather than complete media.

When I compared method #1 and method #2 (both with fixation), the histograms were virtually superimposible +/- 3-4% of 10,000 total events. the second method is now my method of choice and I have found that viability is greatly improved by labeling in complete media. 


Cell Fixation

A dicey area which is always open for debate... "To fix, not to fix, that is the question..."

Sometimes one has to fix the cells for the sake of time. However, the stability of a solution of paraformaldehyde is a concern. Protocols often recommend preparing fresh paraformaldehyde (PF) solutions every month or even every two weeks. Despite this, we have experienced no problems using 1% PF-PBS diluted from a 2% PF-PBS stock that has been stored in the dark at 4 °C for a several months. However, we always observe a very slight shift in the FSC/SSC plots between fixed and unfixed cells for all fixation solutions we have encountered.

In aparticular, we would to mention our experience with a fixative known as FACS/FIX, which was described on the Purdue Flow Listserv. It consists of 13 ml formaldehyde (37% formalin stock), 10 g glucose, and 2.5 ml 10% NaN3, all in PBS to a total volume of 500 ml. This recipe contains methanol as a stabilizer for the formalin. However, we suspected that the methanol would affect light scatter as well as the emission of FITC, PI, or PE. Though easy to make, it had the most dramatic effect on FSC/SSC channel counts we have ever seen.  After fixation in FACS/Fix, the SSC profile was shifted a full log to the right, and the FL1 profile was shifted a half-log to the right. We suspect that either the glucose or the methanol produced this marked shift in the profiles, and we do not recommend this fixative.

David's 2% paraformaldehyde recipe:

For 100 ml final volume. This can be made in as little as 2 hours.
1) Add 2 g paraformaldehye into 50 ml of 2X PBS.
2) Add enough 1N NaOH (~5 ml) to bring the pH to just over pH 8.2, and put it on a rotator or shaking platform at room temperature until parafomaldehyde is dissolved. OR do not add NaOH, and heat to no higher than 60°C while stirring, until the paraformaldehyde is dissolved. 
3) After the PF goes into solution, readjust the pH back to 7.3-7.3 and add water to bring to the solution to a final concentration of 1X PBS.
4) Store foil-wrapped at 4°C.
5) Make up 1% PF-PBS as needed.

Note: Although this is a semantic issue, there is no such thing as a paraformaldehyde solution. Paraformaldehyde is poly-formaldedhyde and is a solid. When para-formaldehyde is in solution, the polyer is broken into the individual formaldehyde monomers.  However, it is common to state one fixes in a certain percentage of parafomaldehyde in PBS as it is so entrenched in contemporary methodology.