Flow Cytometry Core Methods

The protocols we follow in flow cytometry involve cell cycle analysis (checking the cell in its different phases to help detect anomalies), cell labeling (applying antibodies to facilitate antigen detection) and cell fixation (treating cell samples so they do not decay).

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 you are using quickly cycling cells. The easiest cell cycle analysis protocol involves ethanol fixation, staining with PI (propidium iodide) and digestion with RNAse-A. Refer to the protocol from the Current Protocols in Cytometry. 

A Word on the Use of Isotypes
Isotypes are becoming an issue of the past. We are discovering that isotypes can sometimes "lie" to a researcher and render a false positive signal when Fc-receptors are present. These receptors are the most commonly found cells in a myeloid lineage such as macrophages, neutrophils and various granulocytes. We have found that cells that are stressed or are involved, even as bystander cells, in inflammatory processes can be induced to express Fc-receptors (fibroblasts).

Evaluating isotypes must be done with care. If a population is from a tissue, it is likely that there are negative cells for any given marker present. The unstained control still remains the most important control you bring to the flow core.

Isotypes do not account for what is called the "spread of the data" when multiple colors are used in a given experiment.   Fluorescence Minus One, or FMO controls resolve this issue and let us make an accurate determination of the background signal.

For a discussion on the utility of isotypes, you can read Isotype Controls in the Analysis of 

Lymphocytes and CD34+ Stem and Progenitor Cells by Flow Cytometry — Time to Let Go!  We also recommend Flow cytometry controls, instrument setup, and the determination of positivity . In this paper, FMO controls are recommended, especially for dim surface  markers, or markers such as cytokines or FoxP3.  For markers such as these, isotype controls would and do underestimate the background leading the investigator to presume a "false positive" is a real signal.

Cell Labeling

The rule of thumb is to use 1 to 2 μg of primary antibody per 2 to 300,000 cells in a total volume of 100 μl and to keep these ratios consistent between samples and experiments.   Be cognizant of the relative antigen density of your favorite marker.   Always put the brightest fluor on the dimest or least expressed cell surface marker.     Protocol details are as follows.

Method 1

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

This standard protocol has low background, but cells will invariably be lost 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 been 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 LSRII and Fortessa, use 12 X 75 polypropalene snap-top tubes (Fischer #14-9591-0AA). With the tubes labeled, the tops are removed and retained.   The caps 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 LSRII and/or Fortessa. Becton-Dickenson recommends a polystyrene 12 X 75 tube that is only sold through Corning and works well on both machines. However, polystyrene is fairly brittle and if cracked at the lip, it will not work. A solution of 1 to 2 percent 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 label everything in just PBS or you will have very few cells alive before heading to the cytometer.

  1. Re-suspend 3 to 500,000 cells in a final solution of 100 μl of complete media. 
  2. Add 1 to 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 to 7 minutes. 
  4. Add `1 to 2 ug of labeled secondary antibody in 50‒100 μl of complete media and incubate 30 minutes on ice. 
  5. Wash once, adding 4 mL of complete media, centrifuge 500 X g for 5 to 7 minutes. 
  6. Re-suspend in 300 μl of complete media for live analysis; keep on ice and analyze within 1 to 2 hours. If fixation is needed, replace steps 5 and 6 with steps 7 and 8. 
  7. Wash with 4 mL of PBS (no BSA) and centrifuge 500 X g for 5 to 7 minutes. 
  8. Re-suspend in 300 μl of 1 percent paraformaldhye. You can store in the dark and at 4°C for 3 to 4 days.

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

When you compare method 1 and method 2 (both with fixation), the histograms are virtually superimposible, +/- 3 to 4 percent of 10,000 total events. The second method is now the favored method of choice in the lab; viability is greatly improved by labeling in complete media.

Cell Fixation

David's 2 Percent Paraformaldehyde Recipe
For 100 mL final volume; this can be made in as little as two 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 percent PF-PBS as needed.

Although this is a semantic issue, there is no such thing as a paraformaldehyde solution. Paraformaldehyde is poly-formaldedhyde and is a solid. When paraformaldehyde is in solution, the polymer is broken into the individual formaldehyde monomers. However, the common terminology is that you fix a certain percentage of parafomaldehyde in PBS.