Using Flow Cytometry to Detect and Quantitate Altered Blood Formation in the Developing Zebrafish

The Institutional Animal Care and Use Committee (IACUC) advisory board at California State University, Chico, approved all methods described below.

NOTE: It is advised to treat embryos with 1-phenyl 2-thiourea (PTU; Table 1) at 24 hours postfertilization (hpf) to prevent pigmentation which negatively affects fluorescence discrimination by flow cytometry. All procedures listed below are acceptable for flow cytometry and fluorescence-activated cell sorting (FACS). However, if one's goal is to culture the hematopoietic cells after FACS, then adhere to sterile practices when processing samples. The protocol for bleaching and preparing embryo samples in a sterile manner has been described previously¹².

1. Dechorionation of 48 hpf zebrafish embryos

1. With embryos (at least 5, but no more than 200 embryos) in a 10 cm plastic Petri dish, tilt the dish so that the embryos sink to the bottom edge.
   NOTE: It is easy to tilt the dish by removing its lid and placing the lid underneath one edge of the Petri dish.
2. Remove and discard as much E3 medium (Table 1) as possible and add 500 µL of dechorionation protease (10 mg/mL). Incubate at room temperature for 5 min.
3. After 5 min, pick up the Petri dish (keeping all the embryos at the bottom edge of the plate), and gently tap the side of the Petri dish, allowing embryos to gently rub against the bottom of the plate to completely remove chorions.
4. With a squeeze bottle, add ~20 mL of E3 medium to dilute protease. Allow embryos to settle and remove the E3 medium by decanting.
5. Repeat step 1.4 3x to remove all traces of the dechorionation protease.
   NOTE: For each individual sample to analyze, at least 5 embryos are required. This dechorionation procedure can accommodate up to 200 embryos per Petri dish. Samples can be grouped at this stage and separated later if desired. Alternatively, manual dechorionation can be performed by gently tearing the chorions with watchmaker's forceps. This is advantageous when there are small numbers of embryos. Usually zebrafish will hatch from their chorions by 72 hpf. Severely malformed embryos may not, however, and will require dechorionation (either manual or chemical) after that time.

2. Preparation of embryo samples for dissociation

1. Using a P1000 pipette, place 5−10 embryos into a single 1.5 mL microcentrifuge tube.
2. Remove E3 medium with a pipette and discard.
   CAUTION: Pipette with care as embryos can easily be discarded during the following steps.
3. Add 1 mL of 10 mM dithiothreitol (DTT) in E3 medium to remove the mucus coating surrounding the embryonic zebrafish^11,12,13,14. Lay the microcentrifuge tube horizontally, and incubate at room temperature for 30 min.

3. Embryo dissociation

1. Wash the embryos 3x with 1 mL of Dulbecco's phosphate buffered saline (DPBS) with Ca²⁺ and Mg²⁺ to remove traces of DTT. After the last wash add 500 µL of DPBS with Ca²⁺ and Mg²⁺ and 5 µL of 5 mg/mL (26 U/mL) dissociation protease.
   NOTE: Other isotonic buffered solutions may be used, but they must contain Ca²⁺ and Mg²⁺ for the dissociation enzyme to function properly.
2. Incubate samples at 37 °C on a horizontal orbital shaker at 185 rpm for 60 min. Triturate embryos with a P1000 pipette until samples are fully dissociated.
   CAUTION: Take care not to overdigest the embryos; some solid tissue should be present, and it should not be completely homogenous. It is possible to overdigest, which will destroy the target blood cells to be observed on the flow cytometer (Supplemental Figure 1).
   NOTE: This procedure works best for 48−72 hpf embryos. Later stages may require longer incubation with dissociation protease. Timing for different stages must be determined empirically. There are quicker alternative methods to dissociate embryonic tissue, but it is found empirically that this method, although it takes 60 min, is gentle and thorough enough to allow efficient isolation of live hematopoietic cells.

4. Preparation of dissociated embryos for flow cytometry

1. Pipette the 5 dissociated embryos onto the top reservoir of a 5 mL polystyrene round bottom tube with a 35 µmM cell strainer cap.
2. Rinse cells by adding 4 mL of phosphate buffered saline (PBS) with no Ca²⁺ or Mg²⁺ to the strainer cap of the 5 mL polystyrene tube containing the filtered cells.
   NOTE: Other isotonic buffered solutions may also be used, but they should lack Ca²⁺ or Mg²⁺ to dilute and render the dissociation enzyme ineffective.
3. Centrifuge tubes at 4 °C and 300 x g for 5 min to pellet the cells. With a pipette, remove and discard supernatant. Resuspend the cells in 500 µL of PBS (with no Ca²⁺ or Mg²⁺).

5. Flow cytometry

1. Gently vortex the cell suspension and add 1:1,000 red dead cell stain (Table of Materials).
   NOTE: The red dead cell stain is a cell-permeable dye that allows exclusion of dead cells and debris from target cells and is an excellent choice of dye for dead cell discrimination, because it is excited by the 633 nm laser, which is different from the 488 nm laser used to excite common fluorophores such as green fluorescent protein (GFP) and DsRed. In this way, there is no spectral overlap from dead cells and transgenic blood cells. Propidium iodide (PI) at 1 mg/mL is an inexpensive alternative to the red dead cell stain, but make sure to reserve some samples stained only with PI as well as samples only with the fluorophore of interest to set the instrument compensation. Otherwise, the signals of PI and fluorophores such as GFP will overlap, resulting in false-positive results. If the flow cytometer also has a 405 nm laser, other dyes such as blue dad cell stain are also an excellent choice.
   CAUTION: When analyzing blood cells, note that after 30 min certain cell types will begin to phagocytose the red dead cell stain. Keep cells protected from light and on ice until analysis and perform flow cytometry within 30 min of adding the dye.
2. Flow cytometry analysis
   NOTE: Every flow cytometer is a little different, but the following steps will assist in preparing samples for analysis on most machines. For users new to flow cytometry, seek the advice of someone that is an expert on the particular piece of equipment. The following instructions pertain to the BD FACSAria Fusion flow cytometer.
   1. Turn on the cytometer and allow lasers to warm up for 20 min. Empty waste tank, fill sheath tank with appropriate solution (varies based on flow cytometer) and start the fluidics system.
      NOTE: There is usually a particular startup procedure for each type of flow cytometer; follow that startup procedure carefully.
   2. In the analysis software, draw five plots (Figure 1A).
      1. Have the first dot plot measure forward scatter (FSC) on the x axis and side scatter (SSC) on the y axis. Set FSC as linear and SSC as log.
         NOTE: FSC is a measurement of cell size, and SSC is a measurement of granularity; this will allow the discrimination of cells from debris. Embryonic zebrafish blood cell populations do not separate by size and granularity in the same manner as adult zebrafish blood cells described previously¹⁵. Instead they will appear in the same population with all the other digested embryonic cells.
      2. Set the second plot to examine FSC height (H) versus FSC width (W). Set the third plot to measure SSC (H) versus SSC (W).
         NOTE: This step is used to exclude doublets (cells that are stuck together when they pass through the laser).
      3. Set the fourth dot plot to measure the red dead cell stain on the x axis (depending on the cytometer this is usually equivalent to the filter used for allophycocyanin [APC]) and SSC on the y axis. This will allow the discrimination of live cells from dead cells.
         NOTE: Filter sets of each cytometer can be different. Make sure to use the correct detection filter for the dead cell discrimination dye.
      4. The fifth plot measures the fluorophore of choice. Visualize this as a histogram of the fluorophore (Figure 1) or use a dot plot (Supplemental Figure 2).
         NOTE: When examining more than one fluorophore in the same animal, it is recommended to use a dot plot to view both parameters at the same time. If there is any chance that the fluorophores have spectral overlap with each other (Supplemental Figure 2), a dot plot is also recommended. Use the Area (A) settings to calculate the area under the curve generated by the laser pulse of the cytometer for all parameters, because they are more accurate.
   3. Load the sample and reduce the flow rate so that the sample does not run out rapidly. Adjust the FSC and SSC settings so that the bulk of the cell population can be clearly seen (Figure 1A).
   4. Draw a gate around the cell population and label it cells. Making sure that the second dot plot is gated on cells, exclude doublets by first gating on FSC and then on SSC singlets.
   5. On the fourth plot adjust the red dead cell stain settings so that there is clearly a negative population. Draw a gate around these cells, and label them live cells. On the fifth plot, gate on live cells and examine fluorophore negative and positive cells. Draw a gate around the positive cells.
      NOTE: This hierarchical gating strategy examines single fluorophore₊ cells that reside in the live cells gate, which also reside in the cell gate. Take care to set gates properly. These gates can be customized depending on the experiment.
   6. Run each sample, collecting at least 25,000 live cell events.
   7. Follow the shutdown procedure to turn off the fluidics. Refill sheath tank and empty the waste tank.
      NOTE: It is often difficult to visualize fluorophore⁺ cells when the protein expression is low, or the cell population is rare. To ensure the parameters are set properly, fluorophore^– sibling embryos should be prepared alongside to properly draw gates and adjust laser voltages (Supplemental Figure 2). If the target cell population is rare, collect more than 25,000 events. With 5−10 digested animals, one should be able to collect millions of events. The total number of fluorophore⁺ cells can also be obtained with these data. Simply count the total number of cells in the sample with a hemocytometer and multiply the percentage of fluorophore⁺ cells to obtain the absolute number of fluorophore⁺ cells per fish.