Development of the protocol

The study of human DC hematopoiesis has been hampered by the absence of validated markers to identify and track progenitors. Human hematopoietic stem and progenitor cells (HSPCs) are commonly defined by the expression of the cell surface protein CD34, as well as the non-expression of lineage antigens that are present on mature leukocytes. These Lin⁻ (lineage) CD34⁺ HSPCs comprise only a small and variable fraction of human BM cells (2–4%), CB cells (~1%) and peripheral blood (PB; <0.2%). Human HSPCs have been further fractionated using common markers such as CD38, CD90, CD45RA, CD117 (stem cell factor (SCF) receptor) and CD135 (FLT3 receptor)^16-19. However, the combination of these markers does not separate DC lineage–committed progenitors from multipotential progenitors. Moreover, it has been reported that a small fraction of cells that lack the expression of CD34 have progenitor potential²⁰. These Lin⁻ CD34⁻ cells probably represent precursor cells that have lost CD34 expression but that are still too immature to express lineage markers. Therefore, CD34, as well as the afore-mentioned markers, are not enough to further separate HSPCs, and additional markers are required to identify and isolate DC lineage–committed progenitors.

As hematopoiesis relies on instructive cues from the BM niche such as hematopoietins, we hypothesized that the lineage potential of a progenitor might be determined by the set of growth factor receptors that it expresses. We showed that multipotential progenitors such as myeloid and lymphoid progenitors were heterogeneous, and they could be further fractionated using antibodies against receptors for granulocyte–macrophage colony-stimulating factor (GM-CSFR; CD116), macrophage colony-stimulating factor (M-CSFR; CD115) and interleukin-3 (IL-3R; CD123) (refs. 14,15). By using two slightly different antibody panels, we identified the following: (i) the human DC lineage–committed GMDP, MDP and CDP in the Lin⁻ CD34⁺ fraction by combining antibodies against CD45RA, CD116, CD115 and CD123, as well as CD38 (ref. 14), and (ii) the human pre-cDC, which was found in the Lin⁻ CD34⁻ fraction and identified on the basis of the expression or lack of CD45RA, CD116, CD115 and CD123, as well as CD135 and CD117 (ref. 15). These progenitors are rare, which presents a challenge for the isolation of these cells. When working with scarce cells, the simultaneous analysis of all four DC progenitors seems to be essential in order to provide a comprehensive view of DC hematopoiesis.

The proven multicolor panels presented in this protocol are based on our original studies^14,15 with modifications enabling simultaneous assessment of all DC lineage–committed progenitors. As the common feature of progenitors that are capable of developing into DCs is surface expression of CD117 and CD135, the panel to define them includes the following: (i) antibodies specific to CD117 and CD135, as well as CD34, CD123, CD115, CD116 and CD45RA; (ii) antibodies specific to CD3 (T cell), CD19 (B cell), CD14 (monocytes), CD335 (natural killer (NK) cell), CD66b (granulocytes) and CD10 (lymphoid progenitors) to exclude specific lineages (Lin⁻); (iii) antibodies specific to BDCA-1, BDCA-2 and BDCA-3 to exclude differentiated DCs (BDCA⁻); (iv) a viability dye to exclude dead cells; and (v) an optional antibody specific to CD45 to visualize hematopoietic cells in nonhematopoietic tissues or in vitro cultures. All antibodies (Table 1) and fluorochrome combinations (Table 2) are commercially available, and experiments can be run on commonly available flow cytometers and flow sorters that are capable of multicolor analysis (see configurations in Tables 3 and 4).

Defining human DC progenitors also required that we develop an efficient culture assay to study the ability of the progenitors to proliferate and differentiate in response to hematopoietic growth factors. Through systematic testing of different combinations of cytokines, as well as the use of mouse stromal cell lines, we showed that the in vitro differentiation of CD34⁺ HSPCs on MS-5 stromal cells in addition to FLT3L, SCF and GM-CSF cytokines (referred to as MS5+FSG herein) support the development of multiple lineages, including all three major human DC subsets. The resulting DC subsets resemble their blood counterparts both in gene expression and function¹⁴.

Experimental design

In this protocol, we give a flow cytometry–based workflow for (i) simultaneous detection of DC progenitors (Table 2, panel #1 and Fig. 2), as well as their characterization (Table 2, panel #2 and Fig. 3); (ii) isolation of DC progenitors to enable in vitro differentiation (Table 2, panel #3 and Fig. 4); and (iii) evaluation of DC progenitor in vitro differentiation (Table 2, panel #4 and Fig. 5) and proliferation (Table 2, panel #5 and Fig. 6). The procedure also covers various preparatory steps—isolation, freezing and thawing of mononuclear cells (MNCs) and hematopoietic differentiation on stromal cells.

Source tissue

The procedure can be performed on BM or CB samples. BM samples are typically obtained from elderly human donors; however, as age-related developmental changes may affect the composition of the progenitor compartment, CB cells can also be examined. As CB can be collected quite easily and cryopreserved, it is a good source of human progenitors. When these populations are purified and cultured in MS5+FSG, CB progenitors showed a differentiation potential similar to that exhibited by their BM counterparts^14,15.

Identification and characterization of DC progenitors

As described, human DC progenitors can be identified by the lack of lineage and BDCA markers and by combining CD123, CD34, CD115, CD116, CD45RA, CD117 and CD135 markers (Table 2; panel #1). As shown in Figure 2b for BM and Figure 2c for CB, the following cells can be distinguished:

• Cells with the phenotype Lin⁻BDCA⁻CD123^−/loCD34⁺CD117⁺ CD135⁺CD116⁻CD115⁻CD45RA⁺ correspond to the GMDPs with the ability to generate granulocytes, monocytes and DCs.

• Cells with the phenotype Lin⁻BDCA⁻CD123^−/loCD34⁺CD117⁺ CD135⁺CD116⁻ CD115⁺CD45RA⁺ represent the MDPs, which have the potential to give rise to monocytes and DCs but not to granulocytes.

• Cells with the phenotype Lin⁻BDCA⁻CD123^hiCD34⁺CD117⁺C D135⁺CD116⁺CD115⁻CD45RA⁺ correspond to the CDPs with the capacity to give rise to all DCs but not to monocytes and granulocytes.

• Pre-cDCs have the phenotype Lin⁻BDCA⁻CD123^−/loCD34⁻ CD117⁺CD135⁺CD116⁺CD115⁻CD45RA⁺ and are restricted to the two subsets of cDCs.

Of note, we do not include isotype controls in our immunostaining, as the use of fluorescent isotype controls is not completely accurate, and because we have ascertained that the monoclonal antibodies are specific by carefully titrating them and using fluorescence minus one (FMO) controls for gating.

In nonhematopoietic tissues, we recommend using CD45 to visualize hematopoietic progenitors, as it may be challenging to identify these rare cells among other cells such as autofluorescent macrophages. As shown in Figure 2d, most, if not all, DC progenitors express CD45. Therefore, one channel in antibody panel #1 remains open to the potential use of CD45.

As DC progenitors are rare cells in both BM and CB, their identification is a matter of carefully defining the gates separating positive and negative cells. As shown in Figure 2b,c, the first step in the identification of human DC progenitors consists of excluding dead cells and debris by scatter gating. Live cells are then gated for singlets (side scatter(SSC)-W versus SSC-A; W = width, A = area), and dying cells are excluded on the basis of the uptake of the Live/Dead blue stain. The next step consists in the sequential exclusion of all lineage (CD3, CD19, CD14, CD66b, CD335 and CD10)- and BDCA (BDCA-1, BDCA-2 and BDCA-3)-expressing cells. To have a better resolution in our gates, we avoid as much as possible the use of a ‘dump’ channel containing all antibodies marking all excluded populations. Nonetheless, if the use of a ‘dump’ channel is considered to accommodate an additional antibody of interest (as done in Table 2, panel #3), one should carefully titrate each antibody separately before combining them to avoid any compensation issues and difficulties in discriminating between negative cells and cells with low expression. The use of CD123 allows clear separation of cells that express a high level of CD123 (CD123^hi) from cells that express no or a low level of CD123 (CD123^lo cells). CDPs are found within the CD123^hi cells by sequentially gating on CD135⁺, CD116⁺, CD45RA⁺ and CD115⁻ cells. The CD123^lo gate is further analyzed by gating on CD34⁺ CD117⁺ (CD34⁺) cells or CD34⁻ CD117⁺ (CD34⁻) cells. Pre-cDCs are found in the CD34⁻ gate. Indeed, we first gate on cells that express CD135, CD116 and CD45RA but do not express CD115, and then exclude SSC-A-high cells, as these cells contain some monocyte precursors¹⁵. In contrast, the CD34⁺ population contains both GMDPs and MDPs. These progenitors are all CD135⁺, CD116⁻ and CD45RA⁺; expression of CD115 allows the separation of GMDPs (CD115⁻ cells) from MDPs (CD115⁺ cells). Even though the CD115 staining is very faint and it is not possible to get a clear-cut separation of positive and negative cells, the use of FMO control allows for proper gating on CD115⁺ cells (Fig. 2e). Moreover, if we consider that lineage commitment occurs when the capacity to give rise to one cell type, e.g., monocyte and DC, is increased and the capacity to produce another cell type, e.g., granulocyte, is lost, dim CD115 expression is sufficient to separate cells with GMDP potential from cells with MDP potential.

To characterize DC progenitors, we use a related combination of antibodies (Table 2, panel #2). Panel #2 leaves a channel open to accommodate an additional antibody of interest. We prefer the phycoerythrin (PE) channel for this purpose, as PE is bright, and most if not all antibodies are available in this color. Because CD135 staining is crucial in order to identify DC progenitors and because only CD135 PE works well in our hands, we decided to open the PE-Cy7 channel. PE-Cy7 is a tandem dye that can degrade in the presence of light and heat, and it will then emit in the parent dye channel (PE). However, by avoiding the exposure of samples to light and heat, this problem can be largely avoided. The gating procedure is the same as that previously described (Fig. 2b,c), and the expression of CD11c, HLA-DR, CD38 and CD33 on DC progenitors is shown (Fig. 3b). HLA-DR, whose expression is neither specific nor identical within the DC subsets, is also expressed by DC progenitors. CD11c, which is weakly expressed on cDCs and completely lacking on pDCs, is upregulated during DC development. CD38, which was initially used to identify multipotential progenitors¹⁶, as well as DC lineage–committed progenitors¹⁴, is also expressed by pre-cDCs. Because CD38 is expressed by all hematopoietic progenitors, we decided not to use it in our antibody panels and focus more on hematopoietin receptors that allow further separation of these cells. CD33, which is highly specific to the hematopoietic compartment, is expressed by all DC progenitors.

Isolation of progenitors

We developed our flow cytometry strategy to identify DC progenitors on a 16-color flow cytometer (BD LSR II with blue, yellow/green, red, violet and UV lasers; see configuration in Table 3). To isolate DC progenitors, we adapted our panel to maximize the capacities of an 11-color cell sorter (BD Aria III with blue, yellow/green, red and violet lasers; see configuration in Table 4). This was done in part by restricting the number of channels used for lineage markers and by omitting the Live/Dead blue stain to exclude dead cells, relying only on exclusion by scatter gating (Table 2, panel #3). Moreover, the gating hierarchy while sorting is limited to a maximum of an eight-gate sort depth. This limit is intrinsic to the current generation of BD Aria III sorter hardware and firmware. To limit our gating strategy to eight steps (Fig. 4b), we also removed the step to gate out the singlets (SSC-W versus SSC-A). The gating strategy to identify and sort all DC-defined progenitors simultaneously is shown in Figure 4c. The sorted cells are suitable for various downstream applications such as cell culture or gene expression profiling.

Composition and proliferative capacity of in vitro–differentiated cells

Hematopoietic differentiation can be induced in vitro by coculture of hematopoietic progenitors on mouse BM stromal cells plus cytokines. To assess the differentiation potential of different hematopoietic progenitors and their multilineage or restricted potential, we developed an additional antibody panel (Table 2, panel #4). BM CD34⁺ HSPCs cultured in MS5+FSG for 7 d can produce T cells (CD3⁺), B cells (CD20⁺), NK cells (CD335⁺), granulocytes (CD66b⁺), monocytes (CD14⁺) and all three major subsets of DCs, that is, pDCs and both subsets of cDCs (Fig. 5b). Human CD45 antibody must be included in this panel, as it discriminates between human cells and mouse stromal cells, which are not entirely gated out by scatter analysis. Of note, BDCA markers are often used to classify human DCs. However, they are not strictly restricted to DCs; we therefore used them in combination with Clec9A for BDCA-3^hi cDCs, CD14 for BDCA-1⁺ cDCs and CD123 for BDCA-2⁺ pDCs.

To examine the proliferative potential of DC progenitors, we purified them from CB and stained them with carboxy-fluorescein diacetate succinimidyl ester (CFSE) before culturing in MS5+FSG. By using a 14-color antibody panel (Table 2, panel #5) that includes CFSE, we were able to track the cells that were expanding in culture by CFSE dilution. As shown in Figure 6b,c, all progenitors are expanding in culture, even though DC lineage–committed progenitors—GMDP, MDP and CDP cells—have a greater proliferative capacity than DC immediate precursors, i.e., pre-cDCs. Moreover, BDCA-1⁺ and BDCA-3^hi cDCs differentiate as early as 12 h from CDP and pre-cDC progenitors.

Applications of the protocol

We believe that the protocol described here has multiple applications and a wide interdisciplinary scope including (i) further definition of human DC hematopoiesis in health, disease and vaccine settings, as well as (ii) evaluating DC progenitors’ applications in therapeutic approaches.

Our protocol can be used as an initial tool to analyze normal or abnormal immunophenotypic changes during DC development in humans. For instance, it can facilitate the study of diseases involving aberrant DC hematopoiesis such as chronic myelogenous leukemia (CML)²¹ or a newly identified form of primary immunodeficiency characterized by recurrent infections and disseminated bacille Calmette-Guérin (BCG) infection after vaccination in patients lacking GATA2 (ref. 22) and interferon regulatory factor 8 (IRF8)²³. Because of their capacity to elicit and regulate immune responses, DCs are essential to any effort for vaccine development. Therefore, our protocol provides important tools to design and monitor and potentially improve vaccines that rely on DCs.

Finally, the methods described above can support the exploration of new cellular therapeutic approaches based on DC progenitors including stem cell therapies focused on embryonic stem cells and induced pluripotent stem cells. The composition and the properties of the cells that are produced after in vitro differentiation of progenitors will differ considerably depending on the culture condition, and therefore this system can be used to evaluate DC progenitors to identify novel regulators of DC hematopoiesis or to test the toxicity of new drug candidates. Of note, therapeutic approaches based on human DC progenitors may require the development of a nonxenogeneic culture system; therefore, it would be good to explore the feasibility of expanding DC progenitors in an entirely human coculture system using either umbilical vein endothelial cells for CB progenitors or BM stromal cells for BM progenitors.

REAGENTS

EQUIPMENT

• Biological safety level 2 (BSL-2) tissue culture facilities

• BSL-2 sorting facilities

• Class II biological safety cabinet (The Baker Company, cat. no. SG403)

• Cell incubator (37 °C, 5% CO₂; Nuaire, cat. no. NU-5500)

• Tabletop centrifuge with rotors and adapters for 50- and 15-ml conical tubes and 96-well plates (Sorvall, cat. no. 75-004-367)

• Microcentrifuge (Eppendorf, cat. no. 5424)

• Inverted microscope (Nikon, cat. no. Eclipse TS100)

• Hemocytometer (Fisher Scientific, cat. no. 02-671-5) or other cell counter

• Cryo 1 °C freezing container (Nalgene, cat. no. 5100-0001)

• LN2 storage systems (Thermo Scientific, cat. no. 7404)

• BD LSR II flow cytometer running DIVA 8.0, or equivalent (BD Biosciences, special order). Our LSR II is equipped with four lasers with a configuration allowing 16-color, 18-parameter flow cytometry (Table 3)

• BD Aria III cell sorter running DIVA 8.0, or equivalent (BD Biosciences, special order). Our Aria III is equipped with three lasers with a configuration allowing 11-color, 13-parameter flow cytometry (Table 4)

• BD FACSDiva V8.0.1 software (BD Biosciences) for data acquisition

• FlowJo software V9.8.2 (TreeStar) for data analysis

REAGENT SETUP

Donor CB and BM collection and initial processing ● TIMING 10–45 min

! CAUTION All human blood, body fluids and tissues must be treated as if infectious and universal precautions must be taken by lab workers while manipulating these specimens.

! CAUTION All human biological waste to be discarded should be treated with bleach or it should be autoclaved, and then it can be discarded according to your institution disposal procedures.

▲ CRITICAL All steps should be carried out in a class II biological safety cabinet.

• 1|

  Obtain human samples from CB, as described in option A, or obtain and prepare a single-cell suspension of BM, as described in option B.

Isolation of CB and BM MNCs via density gradient ● TIMING 1.5 h

• 2|

  Prepare CB (option A) or BM (option B) density gradients.

Freezing of CB and BM MNCs ● TIMING 20 min

• 9|

  Centrifuge the cells at 480g for 5 min at 20 °C.

• 10|

  Resuspend the cells at a density of 10 × 10⁶ cells per ml in freezing medium.

• 11|

  Transfer 1 ml of cells in freezing medium per sterile cryogenic micro tube.

  ▲ CRITICAL STEP Label each tube clearly (date, sample name, cell concentration).

  ▲ CRITICAL STEP The optimal cell concentration for freezing is 10 × 10⁶ cells per ml. However, you can go up to 50 × 10⁶ cells per ml in case of large specimens.

• 12|

  Put the cryogenic vial in a freezing container, and store it at −80 °C for 12–24 h before transferring it to the liquid nitrogen tank for long-term storage.

  ■ PAUSE POINT Cells can be stored in liquid nitrogen for years.

Thawing CB and BM MNCs ● TIMING 30 min

• 13|

  Add 10 ml of cold thawing medium in a 15-ml conical tube to thaw one vial or add 40 ml of cold thawing medium in a 50-ml conical tube to thaw up to ten vials.

  ▲ CRITICAL STEP To obtain the best possible survival, thawing of cells must be performed as quickly as possible, and the thawing medium should be cold to minimize heat shocks.

• 14|

  Remove the vial from the liquid nitrogen tank, and hold it at room temperature until the sides are thawed but the center remains frozen.

  ▲ CRITICAL STEP Do not place the vial into a 37 °C water bath, as it is important not to dramatically heat-shock the cells.

• 15|

  Transfer the contents of the vial (~1 ml) into a 15-ml conical tube containing cold thawing medium.

• 16|

  Centrifuge the cells at 480g for 5 min at 20 °C.

• 17|

  Resuspend the cells in staining buffer for extemporaneous use.

Culture of mouse MS-5 stromal cells ● TIMING 8 d

▲ CRITICAL Mouse MS-5 stromal cells are only required for staining panel #4 or 5.

• 18|

  Culture MS-5 stromal cells in 10-cm tissue culture dishes with 10 ml of MS-5 growth medium. Place the cells in a CO₂ cell incubator.

  ▲ CRITICAL STEP MS-5 stromal cells grow in monolayers. Examine the tissue culture dish under the microscope; the cells are ready to be split when they are 95% confluent.

• 19|

  Aspirate the MS-5 growth medium and wash the cells twice with 10 ml of PBS.

• 20|

  Add 2 ml of trypsin-EDTA 0.05% (wt/vol) solution and incubate the cells for 2 min at 37 °C in a CO₂ cell incubator.

  ▲ CRITICAL STEP Inadequate washing of cells with PBS or using an old trypsin solution may result in partial detachment of cells from the dish.

• 21|

  Add 10 ml of MS-5 growth medium and collect the cells by pipetting.

• 22|

  Transfer the cell suspension in a 15-ml conical tube and centrifuge it at 480g for 5 min at 20 °C.

• 23|

  Aspirate the supernatant and resuspend the cells in 10 ml of MS-5 growth medium.

• 24|

  Add 0.5 ml of cell suspension to 9.5 ml of MS-5 growth medium, and plate the cells onto a 10-cm tissue culture dish. Place the cells in a CO₂ cell incubator.

• 25|

  When the culture is 95% confluent, if required, split the dish for maintenance (by repeating Steps 19–24). Confluence should be reached after 3–4 d of culture.

• 26|

  To expand the cells for coculture experiment, prepare an extra tissue culture dish by adding 1 ml of cell suspension to 19 ml of MS-5 growth medium into a 15-cm Petri dish.

  ▲ CRITICAL STEP Do not use cells that have a high passage number or that are growing slowly. The doubling time for healthy MS-5 is ~24 h.

  ■ PAUSE POINT It is possible to pause the protocol at this stage by freezing and storing MS-5 stromal cells in liquid nitrogen tank for future use. Cells can be frozen and thawed as described in Steps 9–17 for MNCs.

Hematopoietic differentiation on MS-5 stromal cells ● TIMING 8 d

• 27|

  The day before the coculture experiment (day 0), take the extra tissue culture dish of MS-5 stromal cells that was prepared (Step 26), and proceed to mitomycin C treatment. Aspirate the medium without disturbing the MS-5 stromal cell layer. Add 20 ml of fresh MS-5 growth medium containing 10 μg/ml mitomycin C. Incubate the dish for 3 h in a CO₂ cell incubator.

• 28|

  Trypsinize the MS-5 stromal cell layer, as described in Steps 19–23.

• 29|

  Count the MS-5 cells using a hemocytometer or other cell counters and determine viability by trypan blue stain exclusion.

• 30|

  Centrifuge the cells at 480g for 5 min at 20 °C; aspirate the supernatant.

• 31

  Resuspend the pellet at 2.5 × 10⁵ per ml in MS-5 growth medium.

• 32|

  Seed 2.5 × 10⁴ cells (100 μl) per well of a 96-well flat-bottom tissue culture plate. Incubate the cells overnight in a CO₂ cell incubator.

• 33|

  The following day (day 1), resuspend a maximum of 10⁴ bulk CD34⁺ cells or purified progenitors (see Step 36B for isolation of progenitors using panel #3) in 100 μl of MS-5 growth medium containing 40 ng/ml GM-CSF and SCF ligand and 200 ng/ml FLT3L.

  ▲ CRITICAL STEP If progenitors need to be stained with CFSE (panel #4) before coculture with MS-5 stromal cells, proceed as described in Box 1.

• 34|

  Seed the progenitor cells on top of the MS-5 stromal cells prepared in Step 32. Incubate the cells for 7 d in a CO₂ cell incubator.

Cell surface staining ● TIMING 1–2 h

• 35|

  Take all or an aliquot of the cell suspension from Step 8. Use at least 2–4 × 10⁶ cells for simple phenotyping (panel #1) and characterization (panel #2) staining. If you are sorting the cells (panel #3), start with a sufficient number of cells to obtain a reasonable number of progenitors after sorting. We usually sort 100–200 × 10⁶ CB or BM MNCs to achieve sufficient progenitors. Use the whole MS-5 culture from Step 34 for culture output (panel #4) and culture proliferation (panel #5) staining.

  ▲ CRITICAL STEP We recommend working with fresh samples. If you are working with frozen samples, thawing and staining should preferably be performed on the same day. If you are using frozen cells, count the cells (from Step 17) after thawing using the hemocytometer or other cell counter and determine the viability by Trypan Blue Stain exclusion.

  ? TROUBLESHOOTING

• 36|

  If you are performing cell surface staining for panel #1 and 2, follow option A; for panel #3, follow option B; and for panel #4 and panel #5, follow option C.

  ! CAUTION This and all subsequent steps should avoid exposure to direct light to preserve fluorochromes.

  ▲ CRITICAL STEP Do not forget to prepare the compensation tubes as well (see Reagent Setup).

Acquisition and cell sorting ● TIMING ~1–4 h

• 37|

  Run the compensation controls (see Reagent Setup). Create the compensation matrix and apply it to tubes for both acquisition and sorting.

  ▲ CRITICAL STEP Compensation controls must consist of a negative and a positive population for each single fluorochrome. The negative and positive populations should have the same autofluorescence, which means that each positive bead should be compared with its matching negative bead.

  ▲ CRITICAL STEP When you first set up a panel, you must include a nonstained tube, as well as FMO control (FMOC) tubes. FMOCs sequentially contain all fluorochrome or tandem dyes minus one; they are used to define proper gates and to control the compensation process.

  ? TROUBLESHOOTING

• 38|

  Run the samples (from Step 36A(xii) and Step 36C(iii)) for acquisition on the flow cytometer. Analyze data using FlowJo, as depicted in Figure 2 (panel #1), Figure 3 (panel #2), Figure 5 (panel #4) and Figure 6 (panel #5).

  ▲ CRITICAL STEP If you do not have FlowJo software, use FACSDiva software or an equivalent.

  ? TROUBLESHOOTING

• 39|

  Proceed to simultaneous sorting of the four populations of interest—i.e., GMDPs, MDPs, CDPs and pre-cDCs—from MNCs obtained in Step 8 and stained in Step 36B(i-vi) using an 85-μm nozzle cell sorter. Collect the sorted cells in 1.5-ml conical-bottom tubes with screw caps, in which you add 300 μl of MS-5 growth medium and vortex the collection tubes to coat the surface. You can collect up to 2.5 × 10⁵ cells per tube.

  ▲ CRITICAL STEP If you are sorting cells to analyze mRNA expression by gene array, cells should be recovered without interruption and should be kept at 4 °C at all times to avoid changes in gene expression.

  ? TROUBLESHOOTING

  ? TROUBLESHOOTING

Step 4

If the upper layer is contaminated with RBCs, the samples—that is, upper layer and interphase—should be mixed and layered over a second batch of Ficoll in a new 50-ml conical tube. Repeat this step if necessary.

Step 35

Post-thaw samples may clot. The cells are probably aggregating because of the release of DNA by cells dying during the freezing-thawing process. Include DNase such as benzonase (final concentration 25 U/ml) in thawing medium (Steps 13–17), as well as in staining buffer in Steps 36A(iii,iv), 36A(vii,viii) and 36B(ii,iii).

Step 37

If you have no staining of compensation beads, it may be because not all compensation controls have been prepared in Step 36. You must prepare a compensation control for each fluorochrome or conjugate if you are using tandem dyes.

It may also be that you are not using the proper beads; use anti-mouse Igκ for mouse monoclonal antibodies and anti-rat Igκ for rat monoclonal antibodies.

Step 38

If you have clogging of the sample line while acquiring samples, the cells are probably aggregating because of poor viability. We recommend resuspending the cells in sorting buffer that contains EDTA and filtering the cells using 5-ml polystyrene round-bottom tubes with cell-strainer caps.

Step 39

If you have clogging of the sample line while sorting samples, the cells are probably aggregating because of poor viability. We recommend diluting the cells and filtering the sample again using a 5-ml polystyrene round-bottom tube with a cell-strainer cap.

Staining panel #1: ~3 h–5 h 30 min

Steps 1–8, fresh samples: ~2 h or Steps 13–17, frozen samples: 30 min

Steps 35 and 36A, staining: 1 h 30 min

Steps 37 and 38, acquisition: ~1–2 h

Staining panel #2: ~3 h–5 h 30 min

Steps 1–8, fresh samples: ~2 h or Steps 13–17, frozen samples: 30 min

Steps 35 and 36A, staining: 1 h 30 min

Steps 37 and 38, acquisition: ~1–2 h

Staining panel #3: 3 h 30 min–7 h

Steps 1–8, fresh samples: ~2 h or Steps 13–17, frozen samples: 30 min

Steps 35 and 36B, staining: 1 h

Steps 37 and 39, sorting: ~2–4 h

Staining panel #4: 3 h–5 h 30 min + 8 d coculture

Steps 1–8, fresh samples: ~2 h or Steps 13–17, frozen samples: 30 min

Steps 18–26, culture of mouse MS-5 stromal cells: 8d

Steps 27–34, hematopoietic differentiation on MS-5 stromal cells: 8 d

Steps 35 and 36C, staining: 1 h 30 min

Steps 37 and 38, acquisition: ~1–2 h

Staining panel #5: ~3 h–5 h 30 min + 8 d coculture

Steps 1–8, fresh samples: ~2 h or Steps 13–17, frozen samples: 30 min

Box 1, CFSE staining: 30 min

Steps 18–26, culture of mouse MS-5 stromal cells: 8d

Steps 27–34, hematopoietic differentiation on MS-5 stromal cells: 8 d

Steps 35 and 36C, for staining: 1 h 30 min

Steps 37 and 38, acquisition: ~1–2 h