Isolation of Murine Embryonic Hemogenic Endothelial Cells

Ethics Statement: The protocol outlined below has been reviewed by, and is in compliance with the guidelines of, Yale University's Institutional Animal Care and Use Committee. 

1. Whole Embryo Ex Vivo Culture for Yolk Sac Studies (Optional)

1. Euthanize pregnant dams at E7.0 – E7.5, and remove uterine horns under sterile conditions, as described in greater detail below (steps 2.4 – 2.7).
2. Separate whole embryos (with yolk sac intact¹²) from surrounding decidua, and suspend in 50 ml whole rat serum in 50 ml polystyrene tubes.
3. Gas embryo bottles for 3 min with 5% CO₂ immediately as previously described^12,18. Repeat this step at 24 hr if culturing embryos for 24 – 48 hr.
4. Incubate in rolling 37 °C culture for up to 48 hr.
   Note: Embryos can be treated ex vivo with pharmacological agents (i.e., Notch inhibitor DAPT¹²) or soluble proteins (i.e., fibronectin¹⁹) through pre-incubation of embryos for up to 2 hr in culture medium containing such factors, or through addition of those factors to the rolling culture medium for the entire length of the ex vivo culture period. Gene expression can be manipulated in embryos by pre-incubation of embryos with optimally titered lentivirus for 2 hr¹². Yolk sac vascular and hematopoietic development can be monitored in real time using transgenic reporter mice and optical imaging techniques.

2. Dissection of Yolk Sac (YS) or Aorta-gonad-mesonephros (AGM) from Mouse Embryos

1. Sterilize lab bench by spraying and wiping down all surfaces with 70% ethanol to reduce contamination in subsequent cell cultures. Place an absorbent underpad on lab bench surface.
2. Sterilize surgical instruments with 70% ethanol. Recommended surgical instruments are two #5 straight forceps, and one 8.5 cm straight scissors.
3. If working with embryos from ex vivo culture, carefully remove whole embryos from 50 ml Falcon tubes and proceed immediately to step 2.8. Otherwise, skip this step and proceed to step 2.4.
4. Euthanize pregnant dam at appropriate embryonic age (E8.5 if harvesting YS; E10.5 if harvesting AGM).
   Note: The described technique employs a dual-method euthanasia approach — lethal dosage of a volatile anesthetic followed by mechanical cervical dislocation — to minimize animal pain and distress, and to ensure minimally invasive and humane termination of animal subjects. This combined technique for small rodent euthanasia is recommended by the American Veterinary Medical Association when performed by a trained and proficient investigator (AVMA Guidelines for the Euthanasia of Animals: 2013 Edition).
   1. Anesthetize mouse using a lethal dose of isoflurane (>5% in O₂) for 3 – 5 min. (CAUTION: Isoflurane is a toxic inhalant; use under a chemical fume hood with appropriate respiratory personal protective equipment.)
   2. Following exposure of mice to isoflurane, check at least three signs of anesthetic level — loss of righting reflex, loss of toe pinch reflex, and reduction of respiratory rate — to ensure subjects have achieved a deep anesthetic plane prior to proceeding with mechanical euthanasia.
   3. Cervically dislocate dam to rapidly induce death.
5. Place euthanized dam supine on underpad and liberally spray lower abdomen with 70% ethanol.
6. Make a vertical incision along the midline of the lower abdominal wall. Make additional ~1 inch horizontal incisions extending right and left from the midpoint of the vertical incision, and dissect away the abdominal wall to fully expose right and left uterine horns each containing multiple gestating embryos.
7. Using forceps, hold one of the two uterine horns and use scissors to separate it from the mesometrium. Place dissected uterus on ice in sterile Hank's Balanced Salt Solution (HBSS) in a 60 mm polystyrene tissue culture plate. Repeat for other uterine horn.
8. Under a standard light dissecting microscope, use forceps to pull away muscular layer of uterine sac to reveal the underlying gestational sac and decidua. Isolate YS (Figure 2A) by gently removing the sac from the enclosed embryo. Remove YS tissue from embryo proper at the embryonic origin of vitelline vessels.
9. To isolate the AGM, remove the YS and transect the embryo below the level of the heart and forelimbs and discard the thorax and head region. Next, transect the embryo just below the level of the hindlimbs and remove and discard the tail tissue. Remove the hindlimbs and excess ventral tissue from the remaining section, which contains the AGM (Figure 2B).
10. Pool YS or AGM tissues from multiple embryos and store on ice in HBSS+ (HBSS supplemented with 10% (v/v) fetal bovine serum and 100 U/ml Penicillin, 100 μg/ml Streptomycin, and 0.3 mg/ml L-glutamine) in a clean 1.5 ml tube.

3. Digestion of Primary Tissue into a Single Cell Suspension

1. Centrifuge tube containing harvested YS or AGM tissues for 5 min at 2,000 x g at 4 °C.
2. Remove supernatant and resuspend tissue in 1 ml of either 0.05% (for YS) or 0.2% (for AGM) collagenase type II diluted in HBSS+. Incubate for 30 min in water bath at 37 °C, inverting tube every 5 min to mix.
3. Gently mechanically dissociate tissue by passing sample 10 times through a P1000 pipette. If having difficulty aspirating partially digested tissue, pipette tip bore may be widened by cutting off ~5 mm of tip with a pair of scissors.
4. Centrifuge sample for 5 min at 2,000 x g at 4 °C. Remove supernatant and resuspend in 1 ml ice-cold HBSS+.
5. Pass sample through a 70 μm cell strainer.
6. Count cells using a manual or automated hemocytometer.
7. Centrifuge sample for 5 min at 2,000 x g at 4 °C. Resuspend sample in DMEM+ (4.5 g/L glucose Dulbecco's Modified Eagle Medium supplemented with 10% (v/v) fetal bovine serum and 100 U/ml Penicillin, 100 μg/ml Streptomycin, and 0.3 mg/ml L-glutamine) pre-warmed to 37 °C such that cells are at a final concentration of 1 x 10⁶ cells/ml.

4. Treatment of Cells with Nucleic Acid Dye and Labeling with Fluorescently Conjugated Antibodies

1. Aliquot at least 100 μl of sample (1 x 10⁵ cells) into fresh 1.5 ml tubes for antibody incubation.
2. Include the following sample and necessary control tubes: Unstained; cKit-APC only; CD45-FITC only; CD31-PE only; Flk1-PECy7 only; Hoechst 33342 only; Hoechst 33342 only + Verapamil; Sample (receives all colors, will not receive Verapamil). Use a minimum of 1 x 10⁵ cells in 100 μl for control tubes.
   Note: A larger volume of cells may be added to the Sample tube at the same concentration to maximize yield of sorted Hemogenic EC and HSPC.
3. Add Verapamil diluted in 95% ethanol (see Discussion for rationale) to the “Hoechst 33342 only + Verapamil” control tube to a final concentration of 50 μM. Incubate all tubes for 5 min at 37 °C. (CAUTION: Verapamil is a potent calcium-channel blocking agent and is extremely toxic. Handle with gloves).
4. Add Hoechst 33342 to the “Hoechst 33342 only” control, to the Hoechst 33342 + Verapamil only control, and to the “Sample” tube to a final concentration of 5 μg/ml. Incubate all tubes for 1 hr at 37 °C, protected from light. Gently mix by inverting every 15 min. (CAUTION: Hoechst 33342 is a toxic nuclear dye and should be handled with gloves).
5. Centrifuge all samples for 5 min at 2,000 x g at 4 °C. Remove supernatant and resuspend cell pellets at a concentration of 1 x 10⁵ cells/ml in cold HBSS+.
6. Add fluorescently conjugated antibodies to appropriate antibody-only control tubes, and to the “Sample” tube to a final concentration of 2 μg/ml. Incubate on ice for 30 min, protected from light.
7. Centrifuge all tubes for 5 min at 2,000 x g at 4 °C. Remove supernatant and resuspend cell pellet in 500 μl ice-cold HBSS. Strain samples through mesh filter cap of 5 ml round bottom polystyrene FACS tube and store on ice, protected from light, for immediate FACS.

5. Identification and Isolation of Hemogenic Endothelial Cells and HSPC by FACS

Note: This protocol was optimized using a BD FACSAria 5-laser system equipped with a 100 mw 355 nm UV laser, a 200 mw 405 nm Violet laser, a 200 mw 488 nm Blue laser, a 200 mw 532 nm Green laser, and a 150 mw 637 nm Red laser. Cells were sorted in sterile PBS as sheath fluid and under aseptic conditions, and through a 100 μm nozzle with flow rate set to a sample pressure of 1 such that a maximum of 1,500 – 2,000 events are acquired per second to minimize cell stress.

1. Under these settings described above, use “Unstained” and single color control tubes to optimize FACS cell sorter laser intensities and perform multi-color spectral compensation control according to manufacturer’s instructions. Use forward and side scatter to perform live cell and doublet discrimination from total events (see manufacturer’s instructions for more details).
   Note: This protocol typically results in ~70 – 80% viable cells. If problems are encountered with low cell viability, ensure that tissues are initially dissected quickly in ice cold media, and that all steps except those specifying otherwise are carried out on ice, with gentle pipetting to minimize shearing and cell death (see Discussion for greater detail on preserving cell viability).
2. Use a differential linear-scale plot of Hoechst Red vs. Hoechst Blue fluorescence to identify side population (SP) events (Figure 3A). Use “Hoechst 33342 only + Verapamil” control as a negative control to verify that gates are properly drawn for the sample. SP will appear as a shoulder to the left of the non-SP cells, and will be decreased in the Verapamil-treated control. The non-SP population will be used to identify non-hemogenic EC (Figure 3B).
3. Create additional differential fluorescence plots (log-scale axes) and draw daughter gates of the SP to identify hemogenic endothelial cells (Figure 3C-E).
   Note: Hemogenic endothelial cells are profiled as Flk1+/cKit+/CD45- SP cells (Figure 3D), and HSPC are Flk1-/cKit+/CD45+ SP cells (Figure 3E). Hemogenic endothelial cells can be analyzed in parallel with non-hemogenic endothelial cells, which are identified in this approach as CD31+/CD45- non-SP cells (Figure 3B).
4. Retrieve hemogenic endothelial cell or HSPC fraction onto methylcellulose containing plates for hematopoietic culture (see below), or into other buffers for subsequent processing and analysis.

6. Hematopoietic Differentiation in Culture

1. Add 0.5 ml (135 μl/cm²) methylcellulose-based hematopoietic culture media to desired number of wells of a 24-well tissue culture plate at room temperature. Add 0.5 ml sterile water into unused wells to minimize evaporation. Prepare fresh and keep at room temperature until use.
2. Sort 1 – 10 cells for clonal analysis, or up to 1,000 hemogenic endothelial cells (Flk1+/cKit+/CD45- SP cells) or HSPC (Flk1-/cKit+/CD45+ SP cells) for bulk expansion and differentiation directly into each well of the plate containing methylcellulose. Colony formation is detected in approximately 10 – 20%. 
3. In a sterile tissue culture hood, use a P1000 tip with the tip trimmed to widen its bore, to gently resuspend each well of the seeded methylcellulose media 2 – 3 times, taking care to avoid creation of bubbles. This ensures suspension of the sorted cells into the semisolid methylcellulose for optimal growth.
4. Incubate plate at 37 °C with 5% CO₂ for up to 2 weeks. 
5. Monitor single cell cultures for formation of differentiated hematopoietic cell colonies over time (Figure 4).  Score wells for number and type of differentiated hematopoietic colonies at days 1, 3, 7, and 14 by method(s) outlined below in step 6.7.
   1. Score plates on day 1 to confirm confluency and continued viability of the sorted cells.
   2. By day 3, check for early formation of adherent hemogenic endothelial cell colonies with “cobblestone” morphology (see Goldie et al.¹¹ and Marcelo et al.¹² for representation of day 3 morphology). 
   3. By days 5 – 7, check for formation of rounded HSPC clusters in wells containing sorted hemogenic EC. HSPC should be observed adjacent to flattened hemogenic EC displaying a “cobblestone” endothelial cell morphology.
      Note: Hemogenic endothelial cells should form hematopoietic colonies (determined by hematopoietic colony number), and should demonstrate multi-lineage differentiation capacity (determined by observation of multiple colony types from a single cell).  We have previously observed that ~20% of hemogenic EC or HSPC retrieved by FACS survive to form differentiating and proliferating hematopoietic colonies in methylcellulose¹¹. 
6. To assess colony formation by phase microscopy:
   1. Visualize and count colonies at low magnification under a phase light microscope, and identify BFU-E, CFU-GM, and CFU-GEMM colonies by colony morphology, as described by Goldie et al.¹¹. 
7. To assess colony formation by cell morphology:
   1. Aspirate individual colonies from methylcellulose surface into a P1000 tip with the end trimmed to widen bore. This facilitates aspiration of the viscous methylcellulose medium and ensures successful retrieval of the colony. 
   2. Resuspend picked colonies into 200 ml HBSS and spin onto a glass slide using a cytocentrifuge at 28 x g for 5 min.
   3. Fix slides in 100% methanol for 5 min (CAUTION: Methanol is toxic and should only be used in a chemical hood with appropriate ventilation and personal protective equipment).
   4. Submerge slides in 0.04% Giemsa stain for 20 min (CAUTION: Giemsa stain contains methanol. Use in a chemical hood with appropriate ventilation and personal protective equipment).
   5. Rinse slides in deionized water.
   6. Mount slides with coverslips, and image at high magnification under a standard light microscope, comparing cell morphology to that classically observed in hematopoietic cells from adult bone marrow that are cultured in methylcellulose media. For examples, please see manufacturer’s instructions. 
8. To assess colony formation by cell expression of hematopoietic lineage markers by FACS:
   1. Add 2 ml of HBSS to each well of a 24-well plate containing colonies cultured on 0.5 ml methylcellulose (i.e., dilute commercially-available stock methylcellulose 1:4). Pipette up and down to mix, and transfer to fresh tube(s).
   2. Centrifuge sample at 2,000 x g for 5 min at 4 °C using a standard table-top microcentrifuge.
   3. Remove supernatant and resuspend cell pellet(s) in 1 ml HBSS+, pooling samples if so desired.
   4. Centrifuge sample at 2,000 x g for 5 min at 4 °C.
   5. Remove supernatant and resuspend cell pellet(s) in 400 μl HBSS+.
   6. Aliquot sample into four 1.5 ml tubes as follows: Unstained; B220-FITC only; GR-1-FITC only; Ter119-FITC only.
   7. Add fluorescently-conjugated antibody to appropriate tube to a final concentration of 2 μg/ml. Incubate at 37 °C for 15 min.
   8. Centrifuge sample at 2,000 x g for 5 min at 4 °C. Remove supernatant and resuspend cell pellet in ice-cold 0.5 ml HBSS.
   9. Analyze each sample by FACS for expression of each hematopoietic lineage marker: B220 marks B-cells ²⁰; GR-1 marks myeloid cells ²¹; Ter119 marks erythroid cells ²².
      Note: Additional antibodies targeting other hematopoietic lineage markers may be incorporated into this approach, if so desired.