Immobilization of engineered niche ligands on non-tissue culture plates
1. One day prior to the initiation of culture, combine of Notch antibodies (anti-Notch1 and anti-Notch2 antibodies, each at 10 micrograms/ml), or control antibody (Armenian Hamster IgG Isotype Control Antibody, also at 20 micrograms/ml), together with either recombinant fibronectin fragment (5 micrograms/ml, retronectin, FN-CH-296) or recombinant mouse VCAM-1/CD106 Fc Chimera (5 micrograms/ml) to Ca+Mg+-free PBS.
2. Add approximately 0.25 ml ligand solution per well of a non-tissue culture plastic 48-well plate (for other well formats, see suggested amount per well in Supplementary Table 2). Make sure the meniscus clears the well bottom.
3. Incubate plate overnight at 4°C.
4. On the day of culture initiation, prepare enough serum-free media for the number and types of wells being plated (see Supplementary Tables 1 and 2).
5. Aspirate ligand solution and add excess PBS one well at a time ensuring the well does not dry out. Repeat once.
6. Aspirate PBS from the well and add 0.4 ml of serum-free media per 48-well (reserving additional 0.1 ml volume for addition of cell suspension following sorting, below). (For other well formats, see suggested volumes in Supplementary Table 2).
Isolation of VE-Cad+EPCR+CD61+ hemogenic precursors for engineered culture conditions
*For additional sorting details, refer to our previously published methods article6.
1. Set up timed matings of C57BI6/J7 (CD45.2) mice for generating embryonic tissues of the desired age.
2. Harvest embryos from pregnant females at 9.5 to 11.5 days post coitum (dpc), depending on the stage desired.
3. Dissect the AGM (or for E9.5 embryos, the central portion of the embryo proper containing the para-aortic splanchnopleure/P-Sp) from the embryos on ice-cold PBS with 10% FBS, as previously described7, pooling embryos from litters at the equivalent developmental stage based on somite numbers.
4. Collect the pooled P-Sp/AGM tissues in a 15ml conical tube containing 10 ml PBS with 10% FBS on ice. Gravity settle tissues and gently remove PBS/10% FBS. Add 1ml 0.25% collagenase. Place in a 37°C water bath for 25 min.
5. Add 1 ml of PBS/10% FBS. Pipette about 20-30 times with a 1 ml pipette tip to obtain a single cell suspension. Add an additional 8 ml of PBS/10% FBS and centrifuge at 300 x g for 5 min. Discard the supernatant.
Sorting of VE-Cad+EPCR+CD61+ precursors
1. Prepare the blocking buffer for antibody staining. Add 10 micrograms/ml anti-mouse CD16/CD32 (Fc receptor (FcR) block) and 1 microgram/mL DAPI (1 mg/ml stock in H2O) to 1 ml PBS with 10% FBS.
2. Draw up in a 3 ml syringe and pass through a 0.22-micron syringe filter to sterilize. Re-suspend the cell pellet from the dissociated murine embryonic tissues in 500 microliters blocking buffer and incubate on ice for 5 min.
3. Prepare the antibody mix: Add 10 microgram/ml FcR block to 1 ml PBS with 10% FBS and 10 microliters of each fluorochrome-conjugated antibody (see reagent list): VEcadherin-PECy7, EPCR-PerCP-eFluor710, and CD61-APC. Also prepare a separate antibody mix containing isotype controls antibodies (see reagent list) that will be used to stain a small fraction of AGM cells (~10%) for setting sorting gates. (Note: use a 1:100 final dilution of antibodies unless otherwise indicated based on the manufacturer’s recommendations or according to titrations).
4. Draw antibody mix into a 3 ml syringe and pass through a 0.22-micron syringe filter to sterilize. Add 0.5 ml antibody mix to cell pellet in blocking buffer from above. Incubate on ice for at least 20 minutes.
5. Wash the stained cells: Add 9 ml PBS/10% FBS and centrifuge at 300 x g for 5 min, aspirate, and re-suspend the cell pellet in 0.5 ml PBS/10% FBS.
6. Remove cell clumps by pipetting the cell suspension through a 35-micron cell strainer cap on a 5 ml tube. Place on ice until FACS.
7. Perform fluorescence-activated cell sorting (FACS) by first gating on SSC and FSC (using relatively broad gates for FSC-A, as embryonic hemogenic precursors tend to vary in size, and using FSC-W and SSC-W gates to exclude doublets; for sample gating, see6) and gating on live cells as DAPI negative. Gate on VE-Cadherin positive cells and then gate on the subset that is positive for both EPCR and CD61. Use isotype control and fluor-minus one (FMO) samples to set thresholds for gates. Set the sorting machine to the lowest flow rate of 1.0 to minimize shear stress. Sort cells into a 5ml tube containing cold PBS/10% FBS.
8. After sorting, centrifuge cells at 300 x g for 10 min, aspirate, and re-suspend the cell pellet in serum-free media (enough for 100 microliters/48-well). Re-suspend approximately 1-2 embryo equivalent of sorted cells per 48-well (this may need to be adjusted based on the embryonic stage).
9. Add 100 microliters of cells to each 48-well (from plate prepared for engineered niche culture), for a total volume of 500 microliters/well (or for other well formats, total volume indicated in Supplementary Table 2).
10. Culture cells for 6-7 days in a 37°C tissue culture incubator with 5% C02 before harvesting for phenotypic analysis by flow cytometry and transplantation assays. During culture, colonies of highly motile, semi-adherent, dividing hematopoietic progeny should emerge (see Figure 1 and Supplementary movie), but cells should not be confluent at the time of harvest. To harvest cells, pipette vigorously and scrape wells with pipette tip to remove all cells. Remove 10% of cells for phenotyping, and the remaining 90% for transplantation or other secondary assays (such as CFU or lymphoid assays in secondary OP9 and OP9-Delta co-cultures8). General phenotyping and transplantation assays are described in our previously published protocol.6 To ensure that all cells are harvested, a small volume of PBS/2% FBS can be used in an additional round of vigorous pipetting and added to the rest of the harvested cells.