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The FACS DiVa and FACS Aria can sort into 1.5ml Eppendorf tubes, 12 x 75mm test tubes, 15ml conical centrifuge tubes or 96 well plates. In addition, the FACS Aria will sort into 1ml microtubes. Where the sorted population consitutes from 10% to 99% of the original population, 15ml conical centrifuge tubes should be used. They should be filled with 5ml of culture media. If the sorted population is less than 10% of the original, then the 15ml collection tubes should be filled with 10-13 ml of media or 12 x 75 tubes used with several ml of media. If sorting into 96-well plates, 100ul of media should be placed in each well prior to sorting.
Sorted cells ride in droplets composed of sheath fluid on their way to the sort collection tube. Once the cells have arrived in the collection vessel, they are mixed with the sheath fluid from the droplets and culture media that has been placed in the collection tube. There are 3 choices of sheath fluid that can be used in the FACS DiVa or FACS Aria:
Both Facility supplied sheath fluids are essentially PBS with or without an antifungal/antibacterial preservative agent (Proclin 300). Most cell types tolerate exposure to the sheath fluid preservative and thrive after sorting. Some cells, such as human stem cells and human dendritic cells, do not tolerate exposure and tend to die quickly. In experiments where cells may not tolerate exposure to the sheath fluid preservative, we recommend substituting either Facility supplied preservative-free sheath fluid or 1X PBS. To allow enough set up time to prepare the instrument using 1X PBS, the lab requesting the sort should bring 3-6L for the FACS DiVa or 4-10L for the FACS Aria to the Facility the day before the sort. The amount needed for the sort will depend on the length of time scheduled. Please ask one of the Facility personel for advice on the amount of PBS needed.
Spectral overlap between fluorochromes in multi-color experiments requires the use of fluorescence compensation controls. (See http://www.drmr.com/compensation/index.html for an in depth explanation of compensation.) The proper compensation controls include a negative control (unstained cells are recommended) and one tube each of cells (or beads) stained positively with each of the fluorochromes used in the experiment. The negative control establishes the background fluorescence of the experimental samples and is used to set the baseline PMT (photomultiplier tube) voltages of the instrument. Each of the compensation tubes is subsequently run to establish the spill-over values of each fluorochrome into the other fluorescent channels. It is important that each compensation tube have a population of brightly stained cells (or beads) in order for the spill-over values to be accurately determined. Several vendors sell beads specifically for use as compensation controls. The beads are stained as if they were cells using the same antibodies and fluorochromes that are used in the experiment, producing both a negative and bright positive population for each color. For experiments that cannot spare cells for compensation, do not have enough positive events, or have only low antigen expression, compensation beads are recommended. If beads are not used, then cells expressing high levels of antigen (does not have to be an antigen of interest in the experiment) are stained with a fluorochrome-conjugated antibody that yields brightly stained cells. Because of the necessity to have brightly stained cells at a relatively high frequency (i.e, above 10% of the population) for accurate compensation, it may be necessary to use the same antibody that stains the high density antigen while varying the fluorchrome for each tube (see the following example).
Example: For mouse splenocytes stained with FITC, PE, PerCP, and APC conjugated antibodies, compensation controls should include:
or if using beads
* any antibody that is compatible with the beads
Please Note: The Facility's mission is to serve investigators in their quest to obtain accurate data. The lack of proper compensation controls may yield misleading, confusing, and inaccurate data. In order to live up to the Facility's mission of assuring quality control and reproducibility, Facility staff will not assist with running samples when the necessary compensation controls are not provided by the investigator.
Certain cell types like monocytes, granulocytes, and adherent cell lines tend to be sticky and form aggregates. These aggregates will plug the instrument (a 77um aggregate will not go through a 76um jet). If you can "see" anything in your sample tube, it probably means that the cells have aggregated. Those samples must be filtered with nylon mesh to remove the aggregates or dispersed by some other method before running on the flow cytometer. Add 0.02mg/ml DNase type IIS to all cell preparation steps, including wash steps, to eliminate free DNA from broken cells that leads to aggregation. Cations must be availible to the DNase in order to work properly (i.e., avoid using EDTA). A commercial product, Accumax, has been developed for the specific purpose of keeping cells from clumping. Other sources of large debris such as solid tissue should also be filtered with nylon mesh. In general, anything that you can "see" in the sample tube is too big to go through the instrument.
Just prior to sorting, cells should be filtered through nylon mesh. 70µm mesh filters (Falcon 352350) available through Biochem Stores are recommended. Filtering cells greatly reduces the probability of plugging the instrument during sorting. As a general rule, we will not sort unfiltered samples. We want to insure a successful sort and once the instrument is plugged, it may take as long as an hour to bring the instrument back to its original configuration. Time used to unplug the nozzle and bring the instrument back to sorting status may use up your scheduled time.
Resuspension of cells in media containing phenyl red should be avoided whenever possible. Phenyl red may increase the background fluorescence of cells.
When fixing cells for immunofluorescent experiments with formaldehyde, a common problem is increased autofluorescence. The resultant decrease in separation between the negative and positive populations can render some experiments useless. The most common reason for increased autofluorescence is pH drift of the formaldehyde. It is important that correct pH is established in fresh formaldehyde and that pH is monitored as the fixative solution ages.
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