Quantification of Mouse Hematopoietic Progenitors’ Formation Using Time-lapse Microscopy and Image Analysis

In vitro differentiation of mouse embryonic stem cells (mESCs) towards blood cells constitutes a well-established system to study the endothelial-to-hematopoietic transition (EHT) at the onset of blood development. Assessing the emergence of small non-adherent round blood cells in the culture without disturbing it is essential to evaluate the progression of EHT and also to test conditions potentially enhancing or repressing this process. Here, we describe how to quantify the formation of mouse hematopoietic progenitors during EHT in normal conditions or following over-expression of eight essential transcription factors using time-lapse microscopy and image analysis.


Background
The first hematopoietic stem and progenitor cells (HSPCs) emerge from endothelial cells in the large arteries of the mouse embryo (de Bruijn et al., 2000;Zovein et al., 2008;Chen et al., 2009;). This evolutionarily conserved event is called endothelial to hematopoietic transition (EHT). As a result of EHT, endothelial cells lose their specific markers, start to express hematopoietic genes, gain round morphology and eventually detach from the endothelial layer (Boisset et al., 2010;Kissa and Herbomel, 2010;Rybtsov et al., 2014).
These specific series of events are used to detect EHT activity and monitor the progress of hematopoietic progenitor formation. The widely used method is to quantify endothelial and This article is distributed under the terms of the Creative Commons Attribution License (CC BY 4.0). hematopoietic marker expressing cells. This can be done by gene expression analysis at the single-cell level (Bergiers et al., 2018). Also, protein expression change can be used to assess EHT progression by flow cytometry analysis of endothelial markers such as VE-Cadherin, CD31 and hematopoietic markers such as CD41, CD45, CD43. For spatial information, the same markers can be used for immunofluorescent staining on the fixed tissue. However, all of these techniques require harvesting of the examined cells and therefore represent a single time-point. Besides, the switch between endothelial and hematopoietic gene expressions occurs gradually and in a heterogeneous way so that a marker analysis of a specific time-point is not enough to follow the transition. To complement the gene/protein expression based detection methods, there is a need of a protocol to monitor morphological changes and quantify the round hematopoietic cell progenitors arising through EHT.
To study the hematopoietic cell formation dynamics in vitro, researchers developed embryonic stem cell differentiation protocol (Keller et al., 1993;Kennedy et al., 1997;Sroczynska et al., 2009). The protocol follows the developmental stages of hematopoiesis and recapitulates EHT in vitro (Choi et al., 1998;Nishikawa et al., 1998;Palis et al., 1999). Briefly, mouse embryonic stem cells are cultured to form embryoid bodies containing the in vitro equivalent of hemangioblast, or blast colony forming cells (BL-CFCs), which is then isolated by cell sorting. Those cells grow and give rise to smooth muscle, endothelial and hematopoietic cells (Keller et al., 1993;Faloon et al., 2000). Time-lapse imaging capturing the BL-CFC culture allows us to visualize cells transitioning into hematopoietic progenitors in real-time. Unlike the endothelial and smooth muscle cells, the cells undergoing EHT become round and bud-off from the endothelial cell layer . Here, by combining in vitro time-lapse imaging of an adherent BL-CFC culture with automatic image analysis we introduce a simple and efficient method to quantify those round cells during a culture period, which gives a direct measure of the number of cells undergoing EHT. This protocol enables us to easily test novel parameters affecting EHT rate such as overexpression of certain transcription factors (Bergiers et al., 2018) or testing pathway inhibiting small molecules in the culture media (Vargel et al., 2016). Below, we describe the details of the time-lapse microscopy of BL-CFC culture and image analysis to assess the number of cells underwent EHT.

Procedure
Prior to the start of the BL-CFC culture, ESCs are differentiated according to previously described protocols ). Flk1 + cells are isolated using magnetic sorting following the manufacturer's instructions (Miltenyi Biotech). The procedure described below has been optimized for adherent BL-CFC culture allowing the use of time-lapse microscopy. The quantities and volumes listed were further adapted for the comparison of two culture conditions: with or without doxycycline. Note: Before starting, prepare working solutions as described in "Recipes". Careful, BL-CFC culture medium should be prepared fresh.

1.
Put 1 ml/well of 0.1% gelatin solution (see Recipes) into eight wells of Costar ® 6-well cell culture plates (one plate of four wells for each line) and leave at room temperature (RT) for at least 20 min.

2.
For each line, introduce 0.425 million Flk1 + cells obtained after embryoid body differentiation of ESCs and counted using a Neubauer counting chamber ) into a 15-ml Falcon tube containing 5 ml of IMDM + 20% FBS.

3.
Centrifuge at 290 x g for 5 min.

4.
For each line, remove the supernatant using an aspirating pipet connected to the vacuum pump BVC control, gently tap the bottom of the tube to loosen the cell pellet and resuspend into 8.5 ml of BL-CFC culture medium.
Note: Always use individual pipets and tips for each cell line to avoid crosscontamination.

5.
Aspirate the gelatin from the pre-coated Costar ® 6-well cell culture plate wells using an aspirating pipet.

6.
Resuspend and evenly distribute 2 ml/well of cell suspension into four gelatincoated wells for each cell lines.
Note: This means that 0.1 million Flk1 + cells are added per well.

7.
Agitate the plates parallel to the bench doing vertical and horizontal small jolting movements.
Note: Careful not to do circular movements which would result in the formation of a vortex and, consequently, in all the cells going to the center of the well.

8.
Place the plate into a 37 °C, 5% CO 2 cell culture incubator for 24 h.

9.
Keep the BL-CFC culture medium in the fridge for the doxycycline dilution on the next day.
Note: The differentiation rate can be affected by cell confluence. Make sure the cells are equally spread across the surface of the wells to insure that the emergence of round blood cells will occur at the same rate everywhere in the wells.

1.
Prepare 0.2 μg/μl doxycycline dilution by adding 4 μl of 10 mg/ml doxycycline stock solution (see Recipes) to 196 μl of BL-CFC culture medium kept in the fridge from the day before. Mix well.

2.
Take out the culture plates from the incubator and check under the microscope that the wells are all similar to each other.

3.
Add 10 μl of 0.2 μg/μl doxycycline dilution to each +dox condition well (two wells for each line at a final doxycycline concentration of 1 μg/ml). Add the doxycycline directly in the center of the wells and mix directly after by agitating the culture plates carefully.

4.
Add 10 μl of the BL-CFC culture medium to each -dox condition well as control (two wells for each line).

5.
Put back the two plates inside the incubator and place them carefully into the IncuCyte HD microscope.
Note: Don't write on top of the wells and avoid splashes of the medium on the lid to prevent malfunctioning of the IncuCyte HD as it takes pictures from the top.

6.
While the lid condensation is going away, set up the IncuCyte HD device using the IncuCyte software on the computer:

a.
Connect to the device and log in.

b.
Click on "Schedule Upcoming Scans" in the Task List panel on the left (see Figure 1 A).

c.
In the Drawer Setup window, select an empty vessel on the screen, right-click on the location where you inserted your first plate and click on "New" (see Figure 1).

d.
Keep this vessel selected and select the following settings in the Scan Setup tab, on the left side of the drawer map (see ii. Select the +dox compound, the wells that you treated with doxycycline and click on "Add +dox" (see Figure 4 A).

iii.
Select the concentration and click on OK.

iv.
Click on OK.

g.
Repeat the setup of the vessel (from Day1 Steps 6c-6f) for your second plate.

h.
Right-click on the upper timeline (see Figure 2) and click on "Set intervals".

i.
Set intervals every 15 min starting from time 0, for a total of 24 h.
Note: Even with a total of 24 h, the Incucyte device will scan your vessels every 15 min until you manually remove your vessels from the software.

j.
Check the grey bars to make sure they are not overlapping (see Figure 5 A), check that IncuCyte device is not scanning (see Figure 5 B) and click on "Apply" (see Figure 5 C).
Note: When the plate is put in the incubator, the difference of temperature between the room and the incubator generates water condensation on the lid (as an example, see Figure 6 A). Wait at least 30 min before initiating the imaging so that the microscope can focus properly on the cells.

7.
Come back after 15 min to check that everything is working and switch off the computer if you wish.

1.
After 48-h treatment, click on "Schedule Upcoming Scans" in the Task List panel of the IncuCyte software.

2.
Right-click on the upper timeline and select "Delete Intervals".

3.
Click on "Apply" to stop the scanning by IncuCyte.

4.
Take out your plates and proceed with flow cytometry analysis according to standard protocols.

1.
Export your images from the IncuCyte software:

a.
Click on "Find Scanned Vessels" in the Task List panel of the IncuCyte software (see Figure 6 B).

b.
Select one of your cell line vessels (see Figure 6 C) and click on "View Vessel" button in the lower right-hand corner of the screen (see Figure 6 D).

c.
Check that the scans are fine (no dust, condensation, etc.).

d.
Selecting "Export Movie or Image Set…" from the "Utilities" pull down menu (see Figure 7 A).

e.
In the new window, select the appropriate time frame (see Figure 8 A), the wells corresponding to one condition (see f. Specify the destination folder and the prefix of the files that will be generated (see Figure 8 E).
Note: Create one folder per condition per cell line.

h.
Repeat Steps 1a-1h for all the vessels and conditions to be analyzed (see Supplemental File 1: Example dataset of IncuCyte images for the i8TF cell line for an example dataset).

2.
Analyze your images using CellProfiler software to get the number of round cells for each time-point, cell line and condition: a. Install CellProfiler2.2.0, which can be downloaded from here: http:// cellprofiler.org/previous_releases/ Note: You need Java as well: http://cellprofiler.org/releases/.

b.
Open CellProfiler and load the CellProfiler pipeline (see Supplemental File 2: CellProfiler pipeline for round cell counting) using File > Import > Pipeline from File… in the main menu of CellProfiler.

c.
Drag and drop the whole IncuCyte export destination folder into the File list area of the Images module (see Figure 9 A).

d.
Click on "Analyze images" on the bottom left to start processing (see Figure 9 B).
Note: The pipeline has been developed to automatically generate an output folder named as the input folder with "--analyzed" appended.
The output folder will be generated in the folder containing the input folder. If you want to modify this, simply change the Output File Location to Default Output Folder in the SaveImages and ExportToSpreadsheet Analysis modules, and change the output folder name by selecting "View output settings" in the Output panel of the main window.

e.
Use the Browser to access the output "image.txt" file in the output folder (as an example, see Supplemental f. See Figures 10 and 11, and Supplemental File 3: CellProfiler output files for the example dataset for the analysis output of the example dataset. Notes: i. The Metadata extraction described above relies on the IncuCyte file naming scheme. If this naming scheme is changing, e.g., due to version updates by IncuCyte, or because you are using a different microscope, it will not work. In such cases please contact us and we will help you adapting the Metadata extraction inside CellProfiler. ii. In the output folder, CellProfiler also generates copies of the individual images with all counted round cells marked by a yellow dot (Figure 10).

3.
Generate movies from your images using Fiji software to illustrate differences between round cell emergence rates and highlight other morphological variations:

a.
Install Fiji software, which can be downloaded from here: http://fiji.sc/ Fiji.

b.
Open Fiji.

c.
In your Browser, sort the IncuCyte exported files by Name, select all the files corresponding to the images taken from one frame of one condition for one cell line.
Note: Prepare, aliquot and store at -20 °C. Once thawed, can be stored at 4 °C and used up to one month after.

6.
10 μg/ml VEGF stock solution (stored at -80 °C) Dissolve a 10 μg vial in 1 ml of sterile PBS + 0.1% BSA solution Note: Dissolve, aliquot and store at -80 °C. Once thawed, can be stored at 4 °C and used up to one month after.

7.
10 μg/ml IL6 stock solution (stored at -80 °C) Dissolve a 10 μg vial in 1 ml of sterile PBS + 0.1% BSA solution Note: Dissolve, aliquot and store at -80 °C. Once thawed, can be stored at 4 °C and used up to one month after.

Supplementary Material
Refer to Web version on PubMed Central for supplementary material.