Nuclei Isolation from Mouse Cardiac Progenitor Cells for Epigenome and Gene Expression Profiling at Single-Cell Resolution

The animal procedure adopted in this study was approved by the animal ethics committees of the Aix-Marseille University (C2EA-14) and was carried out according to protocols approved by the appointed national ethical committee for animal experimentation (Ministère de l'Education Nationale, de l'Enseignement Supérieur et de la Recherche; Authorization Apafis N°33927-2021111715507212).

1. Setting up the timed mating prior to dissection

1. To generate mouse embryos, perform a timed mating between adult mice 9.5 days prior to heart region isolation. In this procedure, wild-type C57BL/6J mice aged 2-6 months were used, and timed mating was performed overnight.
2. The next morning, check if the female mice have a vaginal plug. Consider the day the plug is identified as embryonic day (E) 0.5.

2. Tissue preparation and cell isolation

1. Euthanize 2-6 months old pregnant female C57BL/6J at day 9.5 post-conception via CO₂ with an increasing concentration followed by cervical dislocation. Clean the lower part of the abdomen with 70% ethanol.
   NOTE: The use of anesthetics before cervical dislocation is not recommended as this would expose the embryos to environmental changes that could impact subsequent transcriptomic and epigenomic studies.
2. Open the abdomen and peritoneum with scissors. Visualize the uterine horns, and use forceps to excise both horns.
3. Place the horns in a Petri dish containing cold complete medium with 1% FBS. While no specific volume is required, care must be taken to ensure that the embryos are completely immersed in the medium. An approximate volume of 10 mL per 10 cm Petri dish is appropriate.
4. Under the stereomicroscope set at 5x magnification and using forceps, remove the endometrial tissue, placenta, and yolk sac from each embryo.
5. Place the embryos in a Petri dish with cold complete medium with 1% FBS. Dissect the embryonic heart region, as described on the schematic embryo illustrated in Figure 1, in cold complete medium.
6. Pool together the heart regions dissected from five mouse embryos in a 1.5 mL microcentrifuge tube. Prechill swinging bucket centrifuges to 4 °C.
7. Centrifuge at 300x g for 1 min at RT in a swinging bucket centrifuge. Remove the supernatant, and wash the tissues by adding 1 mL of tissue culture-grade PBS.
8. Centrifuge at 300 x g for 1 min at RT. Remove the supernatant, and repeat the washing steps for a total of two washes. Repeat the wash twice, replacing the PBS with 0.05% trypsin/EDTA (1x).
9. For tissue digestion, add 50 µL of 0.25% trypsin/EDTA for 10 min at 37 °C. Apply gentle mechanical dissociation after 5 min by up and down gestures using a wide-orifice filter tip.
10. Inactivate trypsin digestion activity by adding 350 µL of complete medium to the dissociated cells. Pass the resuspended cells through a 40 µm cell strainer.

3. Cell counting and viability assessment

NOTE: The cell count and viability are critical parameters for the success of single-cell experiments. The snATAC-seq approach is sensitive to minor variations in cell number. Too few cells lead to over-digestion of the chromatin, which results in a higher number of reads and the mapping of inaccessible chromatin regions (noise). Similarly, having too many cells generates high-molecular-weight fragments that are hard to sequence. For the accurate determination of the cell and nuclei concentrations, the cell count and viability were assessed by two different methods.

1. Perform a trypan blue assay for cell counting, as described below.
   1. Vortex 0.4% trypan blue staining solution, spin for 10 s at room temperature at 2,000 x g, and transfer 10 µL into a microtube.
   2. Using a pipette, mix the cell suspension, and add 10 µL of suspension to the already aliquoted 10 µL of trypan blue solution. Mix carefully 10 times with a pipette.
   3. Transfer 10 µL of the stained cells into a counting slide. Place the slide into the counter to automatically calculate the cell concentration and viability.
2. Perform a fluorescence-based assessment of mortality as described below.
   NOTE: This assay is based on the use of fluorescent dyes (ethidium homodimer-1, EthD-1, and calcein AM) to evaluate the cell viability. A commercially available kit was used, and the provider's instructions were followed for the appropriate preparation and usage.
   1. Prepare a 10 µM EthD-1 solution by adding 5 µL of the supplied 2 mM EthD-1 stock solution to 1 mL of sterile tissue-culture grade D-PBS, and vortex for 5 s to ensure complete mixing.
   2. Transfer 2.5 µL of the supplied 4 mM calcein AM stock solution to the already prepared EthD-1 solution. Vortex for 5 s to ensure complete mixing.
   3. Add 10 µL of the resulting 10 µM EthD-1 and 10 µM calcein AM working solution to 10 µL of cell suspension.
      NOTE: Calcein is highly susceptible to hydrolysis in aqueous solutions, so use this working solution within 1 day.
   4. Transfer 10 µL of the stained cells onto a counting slide. Insert the slide into the automated fluorescent cell counter. Count the live cells using a GFP filter and the dead cells with an RFP filter.
      NOTE: The two counting methods gave similar cell counts and viability, and the total number of freshly dissociated cells was estimated to average around 20,000 cells per embryonic heart region (0.06 mm³).

4. Cell freezing

1. Centrifuge the cell suspension at 500 x g for 5 min at 4 °C. Carefully remove the supernatant without touching the bottom of the tube, and resuspend the pellet in 1 mL of chilled freezing medium.
2. Transfer the cell suspension into a pre-cooled cryovial, and place the cryovial in a pre-cooled freezing container.
3. Place the container at −80 ^◦C for 4 h to 8 h. Transfer the cryovial to liquid nitrogen for downstream single-nucleus RNA and ATAC sequencing experiments.
   ​NOTE: The cryovial should be transported on dry ice prior to usage. In our experience, the cells may be stored in liquid nitrogen for at least 6 months with no loss of cell viability. The storage temperature should be kept below −150 °C all times to prevent the formation of ice crystals.

5. Cell thawing and removal of dead cells

1. Place the cryovials in a 37 °C water bath for 1-2 min. Remove it when a tiny crystal remains in the cryovial.
2. Add 1 mL of pre-warmed (37 °C) complete medium. Mix with a pipette three times. Transfer the suspension to a 15 mL conical tube containing 10 mL of warm complete medium.
3. Pass the entire cell suspension over a 30 µm strainer. Rinse the strainer with 1-2 mL of warm complete medium, and collect the flowthrough in the same conical tube.
4. Centrifuge at 300 x g for 5 min, and remove supernatant. Wash one time with PBS, and transfer the cell suspension into a 1.5 mL microtube.
5. Centrifuge at 300 x g for 5 min, and remove the supernatant without disrupting the cell pellet.
6. Add 100 µL of the provided magnetic microbeads to the pelleted cells, and homogenize five times using a wide-bore pipette tip. Incubate for 15 min at RT.
7. In the meantime, rinse the magnetic separation column (capacity: 1 x 10⁷ to 2 x 10⁸ cells) with 500 µL of 1x binding buffer.
8. Once the incubation is completed, dilute the cell suspension containing the microbeads with 500 µL of 1x binding buffer.
9. Apply the cell suspension (0.6 µL) to the prepared magnetic separation column. The flow-through is the negatively selected live cell fraction, and the magnetically retained fraction is the positively selected dead cells.
10. Collect the live cell effluent in a 15 mL centrifuge tube. Rinse the 1.5 mL microtube that contained the cell suspension and the column with 2 mL of 1x binding buffer, and collect the effluent in the same 15 mL tube.
11. Centrifuge at 300 x g for 5 min. Remove the supernatant without disrupting the cell pellet.
12. Add 1 mL of the PBS-BSA solution, and mix gently by pipetting five times using a wide-orifice pipette tip. Transfer the cell suspension into a 1.5 mL microtube.
13. Centrifuge at 300 x g for 5 min. Remove the supernatant without disrupting the cell pellet.
14. Add 1 mL of PBS-BSA to resuspend the pellet, and repeat the washing step for a total of two washes.
15. Resuspend the cells in 100 µL of PBS-BSA solution. Mix gently by pipetting 10 times.
16. Determine the concentration and viability of the cells using the previously described methods (step 3.1 and step 3.2). To proceed to the next step, ensure a cell count of ≥100,000 cells. See Figure 2 for representative results.
    ​NOTE: Cryopreservation results in a loss of approximately 40% of the initial starting material of live cells. Depending on the starting number of cells, bead separation on a magnetic column results in high cell viability but divides the total number of cells by two-fold to five-fold. The utilization of a column-based magnetic kit to remove the dead cells is advised only when sufficient starting material is available.

6. Nuclei isolation

NOTE: snATAC and snRNA-seq combined are performed with a suspension of clean and intact nuclei. The optimization of the lysis conditions (lysis time and NP40 concentration) for the used cell type is recommended. The optimal lysis timepoint and detergent concentration are those that result in the maximum number of cells being lysed without disrupting the nuclear morphology. The loss of cells/nuclei can be reduced by using a swinging bucket centrifuge instead of a fixed-angle centrifuge. To minimize nuclei retention on plastics, it is recommended to use low-retention pipette tips and centrifuge tubes. This can increase the nuclei recovery. The coating of the pipette tips and centrifuge tubes with 5% BSA is a less expensive but more time-consuming alternative.

1. Clean the working place and materials with 70% ethanol and RNase removing solution.
2. Prechill swinging bucket centrifuges to 4 °C. Freshly prepare lysis buffer, lysis dilution buffer, 0.1x lysis buffer, and wash buffer (see Table 1). Maintain the buffers on ice.
3. Centrifuge the cell suspension at 500 x g for 5min at 4 °C in a swinging bucket centrifuge. Gently remove all the supernatant without touching the bottom of the tube to avoid dislodging the cell pellet.
4. Add 100 µL of chilled 0.1x lysis buffer. Gently mix by pipetting 10 times. Incubate for 5 min on ice.
5. Add 1 mL of chilled wash buffer, and gently mix by pipetting five times. Centrifuge at 500 x g for 5 min at 4 °C. Remove the supernatant without disrupting the nuclei pellet.
6. Repeat the washes with chilled wash buffer for a total of three washes. Prepare the diluted nuclei buffer. After resuspension, keep the nuclei stock solution on ice.

7. Quality and quantity assessments of the isolated nuclei

NOTE: Based on the initial cell count and estimating approximately 50% nuclei loss during cell lysis, resuspend the appropriate number of cells in cold diluted nuclei buffer. The calculation of the resuspension volume with chilled diluted nuclei buffer is based on the targeted nuclei recovery and the corresponding nuclei stock concentrations recommended by the manufacturer's user guide. See an example of this calculation in Supplementary Protocol 1.

1. Based on the initial cell count and estimating approximately 50% nuclei loss during cell lysis, resuspend the appropriate number of cells in cold diluted nuclei buffer.
2. Determine the actual nuclei concentration. Mix 2 µL of nuclei stock solution with 8 µL of diluted nuclei buffer and add the mix to the pre-aliquoted 10 µL of trypan blue or EthD-1/calceinAM working solution. Count the nuclei using an automated cell counter. Less than 5% of the cells should be viable. See Figure 3 for representative results.
3. Calculate the volume of nuclei stock and the volume of diluted nuclei buffer to obtain a total volume of 5 µL at the recommended concentration for the transposition reaction (refer to the manufacturer's user guide). Proceed immediately to the transposition reaction according to the user guide¹⁷.
   ​NOTE: All the steps from the transposition reaction to the construction of the ATAC and GEX libraries were performed according to the user guide of the kit used for snRNA-seq and snATAC-seq combined¹⁷.

8. Quality analysis of the snRNA-seq and snATAC-seq libraries

1. Before proceeding to next-generation sequencing, validate the quality and size distribution of the final snATAC-seq and gene expression GEX libraries. Assess the quality and quantity of sequences identified in the libraries using fragment analysis systems. See Figure 4 for representative quality assessment results.
   NOTE: The final trace of the snATAC-seq libraries should show the periodicity of nucleosome winding, and the sizes should range from 200 bp to several Kbp, whereas the sizes in the gene expression library should range from 300 bp to 600 bp.