Flow Cytometry Protocol

Prepare your cell suspensions for Flow Cytometry

Although most flow cytometry experiments involve labeling populations of cells that are relatively abundant, the number of cells required will vary depending upon the rarity of your cells. In general, researchers will stain between 1 x 10⁵ and 1 x 10⁶ cells per sample. However, with exceeding rare cell populations like stem cell subsets or specific circulating cell types, obtaining an adequate number of events to interpret your data may require 1 x 10⁹ or greater cell counts. Ideally, with these or any samples, you want to analyze a minimum of 100 events in order to keep the COV of like samples under 10%. If you’re staining a new population of cells, we are here to help. Contact our experienced technical support team for staining advice for your samples.

1. Obtain cell suspensions.
   1. For non-adherent cell populations, wash cells (resuspend in buffer, centrifuge at 400 x g for 5 minutes, aspirate buffer, and resuspend in an appropriate volume of fresh buffer) in flow cytometry staining buffer, resuspend and resuspend in a small volume of buffer.
   2. For adherent cell populations, wash cells (similar to a media exchange) in flow cytometry staining buffer and harvest cells by gently scraping the dish, plate, or culture flask. Avoid trypsin if possible as it may damage cell surface proteins. Collagenase or similar may be used if scraping is not sufficient for recovering adherent cells. Immediately wash cells (as described in 1a) again and resuspend in a small amount of flow cytometry staining buffer.
   3. For tissue samples, obtain a cell suspension homogenizing tissue in staining buffer by pressing the sample through a fine mesh sieve (nylon mesh) using a clean syringe plunger from a 3cc syringe, or similar instrument. This procedure will provide sufficient homogenization for most tissues, but other enzymatic methods are available for difficult samples. For example enzymatic tissue digestion protocols, you can see this protocol for harvesting adipocytes.
2. Count Cells.
   1. Gently mix cell suspension to ensure a homogenous mixture and reserve 20-100 ul to count cells.
   2. Use a trypan blue exclusion stain or similar to exclude dead cells. Obtain cell counts using a hemocytometer or automated cell counter.
3. Resuspend cells to an appropriate concentration in flow cytometry staining buffer. 1 x 10⁶ cells is commonly used for antibody labeling of most cell types.
4. We recommend creating multiple aliquots of cells for each sample in order to obtain measurements in duplicate.

Cell Surface Target Antibody Staining for Flow Cytometry

Fc receptors on many immune cells may bind antibodies and create false positive signals. Anti-CD16 + anti-CD32 antibodies are commonly employed as an Fc-block and may be used to reduce or eliminate this source of noise. For cell populations with high numbers of macrophage, dendritic cells, granulocytes, or other immune cell populations, we recommend the use of a blocking reagent.

1. Based on the number of cells you will require (see the above section), place the appropriate volume of cells suspended in flow cytometry staining buffer into 96 well plates or tubes compatible with your flow cytometer.
2. Add your primary antibody (preferably directly conjugated) to each sample. The dilution of antibody should be experimentally determined by running a pilot experiment on several samples. Too much antibody creates high background fluorescence and too little results in dim positive cells. An optimal antibody concentration can be found by comparing staining index at multiple concentrations and selecting the conditions that result in highest signal to noise. In general, our antibodies are provided at concentrations that are optimized for staining common samples like PBMCs (or samples otherwise specified on the product’s datasheet). Staining should still be piloted to account for variations in experimental conditions and sample types. In general, between 0.25 and 2 ug/ml of antibody is provided and will be listed on the datasheet. If using an antibody that is not provided in a test format, begin optimization steps in this range.
3. Multiple primary antibodies may be added at the same time in this labeling step for multicolor experiments.
   1. Be sure to stain appropriate controls as well. In addition to blank controls and isotype controls, this may also include single antibody controls (or antibody binding beads) and FMO controls (all antibodies but one) to enable compensation for overlapping fluorophores.
4. Incubate on ice for 30 min to 1 hour in the dark (ie. in a drawer in your lab).
5. Wash 1-3 times by resuspending cells in a larger volume of staining buffer (250-300 ul for plates, 1-2ml for tubes), gently mixing, centrifuging samples at 400 x g for 5 minutes and aspirating supernatant.
6. Resuspend in an appropriate volume of staining buffer. Higher concentrations of cells may yield faster data acquisition on the flow cytometer but may also lead to formation of cell aggregates which should be avoided.
7. Proceed to running samples on the flow cytometer.

Intracellular Target Antibody Staining for Flow Cytometry

Staining of intracellular targets is trickier than cell surface antibody labeling. However, with proper fixatives and appropriate permeabilization buffers, you can obtain great data that allows for very precise identification of cells. This is not suitable for cell sorting. If both intracellular and extracellular targets are to be labeled, perform cell surface staining prior to this protocol.

1. If working with secreted proteins, treatment with Brefeldin A or Monesin for at least 4 and not more than 24 hours is required prior to staining.
2. Fix the cells.
   1. Incubate in a small volume (100 ul) of fixative solution for 15 minutes. Fixatives used may include 2-4% paraformaldehyde, or methanol. These reagents cross-link proteins to preserve their location and may impact antibody binding. If unsure of what to use, pilot several options prior to staining experimental samples or email us at technical@enquirebio.com.
3. Permeabilize the cells in a small volume (e.g. 100 ul) of permeabilization buffer for 15 minutes at room temperature.
   1. Staining of cytoplasmic targets is often more easily achieved by permeabilizing cells with a mild detergent solution such as PBS and 0.5% tween-20 or saponin. Use of these buffers requires their continued presence at a low concentration in all remaining stages of the assay (0.1%).
   2. For nuclear targets, other permeabilization buffers like Triton X-100 are better suited for staining. These buffers generally provide more permanent permeabilization of cells. Nuclear staining may be performed at the same time as cytoplasmic staining by simply using one of these buffers, however, if difficulty labeling cytoplasmic targets occurs, it may also be performed after staining cytoplasmic targets in the presence of a more gentle detergent.
4. Wash cells by adding 2ml of staining buffer (plus 0.1% permeabilization buffer if required) and then immediately centrifuging at 400 x g for 5 minutes.
5. Aspirate the supernatant and resuspend in staining buffer at an appropriate concentration for antibody labeling (e.g. 1 x 10 ⁶ cells per ml).
6. As in the cell surface staining protocol, optimize the concentration of each antibody to be used in staining.
7. Add each primary antibody to each sample, making sure to stain controls as appropriate (isotype, compensation, FMO, etc.)
8. Incubate on ice for 30-60 minutes in the dark.
9. Wash 1-3 times as described throughout this protocol.
10. Resuspend cells in an appropriate volume of staining buffer, with care to avoid concentrations that will result in formation of cell aggregates.
11. Proceed to running samples on the flow cytometer.

Running Samples on the Flow Cytometer

1. Run control samples in your flow cytometer to adjust PMT voltages. Adjusting PMT voltages essentially changes the sensitivity of the detectors on your instrument. Higher values will shift events further in the positive direction on any plot’s axis. Generally this is accomplished by adjusting voltages until the negative events are just off the axes (but not condensed up against the axes) while maintaining good dispersion of positive events (ie. not over the instrument’s limit of detection).
2. Experimental samples should also be run to set voltages for forward scatter (FSC) and side scatter (SSC) detectors. These values provide an additional dimension for distinguishing amongst various cell types. Again, the goal is to adjust the voltages to achieve good dispersion which ideally will allow separation of different cells into relatively tight groupings based upon size (FSC) and internal complexity (SSC).
3. Set number of events to collect.
   1. Events may be based upon total events or events in a particular population. If setting events based upon a particular population, the gating scheme must be set up in the instrument prior to data acquisition. Otherwise, gating may be adjusted in later data analysis.
4. Set flow rates (faster is generally… faster, but less accurate). Take into account the number of total events that must be analyzed to achieve adequate analysis of the cells of interest.
5. Collect data from controls and experimental samples (gating and compensation will be largely based upon control samples and may be done after the run during data analysis).
6. Preform an instrument decontamination and clean prior to shutdown.
7. Analyze data using flow cytometry data analysis software such as FlowJo. This software generally automates tricky procedures such as compensation so long as appropriate controls were run.