Single-Cell RNA Sequencing Protocol

Preparation of Collection Plates

Cleaning of the Work space and Preparations

1. Clean working area with 70% (v/v) ethanol and DNA-Off wipes.
2. Thaw aliquots of the reagents (single-cell lysis buffer) on ice or at 4°C.
3. Cool metal stands on ice or at 4°C.
4. Once all reagents are thawed, mix them vigorously.

Place Single-Cell Lysis Solution in Plates

1. Prepare the single-cell lysis buffer in a microfuge tube. Mix it by inverting the tube at least ten times, and collect at the bottom by a quick spin in a microfuge (5-10 sec at less than 1000 g). Avoid vortexing the single-cell lysis solution because the detergent will foam up and sequester cells unnecessarily.
2. Assemble the PCR frame plates (four 24-well plates per frame for a 96-well frame plate), and place them onto a nice-cold metal stand to cool.
3. Transfer 3 μL of the single-cell lysis buffer into each well using a multichannel pipette.
4. Seal the PCR plates. Collect the lysis solution at the bottom of the wells by centrifugation for 15 sec at <2000 g.
5. Keep the plates on ice for use within the day.

Dissociation

Preparations

1. One hour before starting the dissociation, supplement the cell culture with 5 μM Y-27632 ROCK inhibitor. It is included in all subsequent steps to prevent apoptosis during dissociation and processing.
2. Prepare sufficient FACS buffer, working dilution of TO-PRO-3 iodide, and tip blocking solution.
3. Warm up TrypLE Express supplemented with 5 μM Y-27632 ROCK inhibitor.

Dissociation and Staining

1. Gently remove the media from the cells cultured in 12-well plates with a micropipette.
2. Gently and without agitation, wash with 2 mL HBSS.
3. Add 0.5 mL TrypLE Express supplemented with 5 μM Y-27632 ROCK inhibitor per well.
4. Incubate at 37°C on an orbital shaker at ≤100 rpm. After 20 min gently pipette up and down the cells 10-12 times, followed by further incubation with agitation. Repeat the pipetting every 5 min until single-cell suspension forms.
5. In the meanwhile, add TO-PRO-3 iodide to the FACS buffer to 1 μM final concentration (except for the unstained control).
6. Collect cells and spin down at 200g for 4 min.
7. For the first sample, also include the ethanol-fixed cells and spin them down too.
8. Resuspend in 300 μl FACS buffer and filter through a FACS filter mesh cap.
9. Maintain the single-cell suspension in FACS buffer on ice.
10. Proceed to FACS the cells within 30 min.

Stained Dead Cell Control Sample

1. Prepare fresh 70% ethanol with absolute ethanol and distilled water. Transfer 3 ml to a FACS tube and chill it on ice.
2. Dissociate cells with TrypLE Express as above.
3. Resuspend in 5 ml medium and spin at 200 g for 4 min.
4. Collect cells in 500 μl HBSS.
5. Drip the cell suspension slowly into the ice-cold 70% ethanol while vortexing the FACS tube vigorously to avoid cell aggregates. Do not allow drops of cell suspension to fall onto the plastic directly.
6. Leave the samples on ice for at least 1 h to permeabilize.
7. On the day of the single-cell collection, spin down, and resuspend the fixed and permeabilized cells in 300 μL FACS buffer with TO-PRO-3 iodide.

Fluorescence-Assisted Cell Sorting (FACS) of Live Cells

Preparations and Flow Cytometer Settings

1. Ice box with the prepared collection plates.
2. Dry ice box with chilled metal stand for snap-freezing the single-cell lysates.
3. Centrifuge to spin down the collection plates.
4. Adhesive seals.
5. Ice box with the samples.

Sorting Strategy

1. From allevents: In FSC-H×SSC-H, select cells of the appropriate size and granularity (gateP1) to exclude debris.
2. From gateP1: In FSC-A×FSC-W, select singlets (gateP2).
3. From gateP2: In SSC-A×SSC-W, select singlets again (gateP3).
4. From gateP3: In FSC-H×TO-PRO-3 iodide, separate live soma (gateP4) from dead soma, apoptotic bodies, and other large debris.

Collection

1. From gateP4 (live soma), index sort one event (cell) per well, and dispense 5-nL volume into 3 μL lysis buffer.
2. After finishing collecting a plate, put it into an ice-cold metal stand within the laminar flow unit, and seal it immediately with sealing foil.
3. Collect lysates at the bottom of the wells by centrifugation for 15 sec at <2000 g.
4. Immediately place the plate onto a metal stand cooled on dry ice to snap-freeze the single-cell lysates.
5. Place the plates into a cooled zip-lock bag on dry ice.
6. Store at -80°C until processing the lysates.

Single-Cell Smart-Seq2 cDNA Libraries

Preparations

1. Clean working area with 70% ethanol and wipes.
2. Thaw aliquots of reagents on ice.
3. Cool metal racks on ice.
4. Once all reagents are thawed, mix vigorously.

First Strand Synthesis

1. Prepare the first strand synthesis reaction mix. Invert the tube, and collect by a quick spin in a microfuge.
2. Set the thermos cycler to 72°C.
3. Place the frozen lysates onto an ice-cold metal stand. Seal the plastic cover shut using a scraper.
4. Centrifugate for 15 sec at <2000 g, and collect the cell lysates at the bottom.
5. Remove the PCR tubes trips from the 96-well framed plate. The strips are held together by the plastic seal. Seal tightly and cut off the excess border of the plastic seal.
6. Place the tubes trips in a thermocycler, put the cover on, and incubate at 72°C for 3 min. This is the denaturation step of reverse transcription.
7. Put the PCR tubes trips back into the 96-well framed plate. Immediately cool on ice for at least 2 min.
8. While the samples are cooling, start up the thermocycler program "1_RT_10" to warm up to 42°C.
9. Collect the samples by brief centrifugation. Remove seal carefully, and place Piko PCR framed plate onto an ice-cold metal stand.
10. Distribute the first strand reaction mix into an eight-well strip (36.5 μL per well). Add 2.76 μL first-strand reaction mix to the side of each well with single-cell lysate using a multichannel pipette.
11. Seal the PCR framed plate.
12. Combine the first strand synthesis reaction with the lysates by centrifugation for 15 sec at <2000 g.
13. Seal the plate with a plastic cover.
14. Remove PCR tubes trips from the 96-well framed plate. The strips are held together by the plastic seal.
15. Seal tightly and cut off the excess border of the plastic seal.
16. Place the plate into the thermocycler, and add cover.
17. Run the "1_RT_10" program.

Second Strand Synthesis and Amplification

1. Prepare the second strand synthesis reaction mix on ice in a microfuge tube. Mix the second strand synthesis reaction mix by inversion, and collect by centrifugation for 15 sec at <2000 g.
2. Distribute the second strand reaction mix into an eight-well strip (100 μL per well) as a reservoir for pipetting.
3. Remove the first strand synthesis reaction samples from the thermocycler. Immediately place the Piko PCR tube strips back into the 96-well plate frame and onto an ice-cold metal stand.
4. Start up the thermocycler program "2_ISP_21".
5. Seal with a plastic cover sheet, and collect reactions by brief centrifugation for 15 sec at <2000 g. Place the Piko PCR framed plate onto an ice-cold metal stand, and remove the seal carefully.
6. Add 7.5 μL second strand reaction mix to the side of each well with the samples using a multichannel pipette.
7. Seal the PCR framed plate. Combine the reaction by brief centrifugation for 15 sec at <2000 g.
8. Remove PCR tube strips from the framed plate. The strips are held together by the plastic seal. Seal tightly and cut off the excess border of the plastic seal.
9. Place the second strand reaction tubes into the thermocycler, add cover, and run the "2_ISP_21" program (continue to the second step). This is the second strand synthesis and amplification step.

cDNA Library Purification

1. Equilibrate the SPRI beads (19.5% PEG-8k) to room temperature for at least 20 min.
2. Distribute the magnetic beads into an eight-well strip (220 μL per well).
3. Column by column, add 12.5 μL beads per well to a 96-well deep well plate using a multichannel pipette.
4. Remove the samples from the thermocycler. Place the Piko PCR tube strips into the 96-well plate frame and onto an ice-cold metal stand.
5. Seal the plate with a plastic cover, and collect reaction by brief centrifugation for 15 sec at <2000 g. Remove seal carefully, and place the 96-well framed plate onto an ice-cold metal stand.
6. Column by column using a multichannel pipette, transfer the samples to the deep well plates with the magnetic beads. After each transfer, mix beads and reaction by pipetting up and down 10 times.
7. After the last transfer, cover the samples, and incubate at room temperature for 8 min to allow the magnetic beads to bind the samples.
8. Place the covered plate with the samples onto a magnetic stand to collect the beads for 5 min.
9. Discard the supernatant without disturbing the beads. Dry the beads for 10 min at room temperature without cover while the plate is still on the magnetic stand.
10. Distribute elution solution into an eight-well strip (160 μL per well) as reservoir for the multichannel pipette. While the samples are still on the magnetic stand, add 13 μL elution solution per well.
11. Remove the plate from the magnetic stand. Column by column, resuspend the beads in the elution solution by pipetting up and down 10 times.
12. Cover the deep well plate, and incubate it at room temperature for 5 min to release the samples from the beads.
13. Place the deep well plate onto the magnetic stand and bind beads for > 2 min.
14. Label 12 low nucleic acid-binding eight-well strips, and place the min to an ice-cold metal stand.
15. Column by column, transfer 12 μL of the eluate from the plate on the magnetic stand to eight-well strips.
While transferring, do not disturb the beads. After the transfer and taking samples for quantification, seal the eight-well strips, and freeze at -20°C.

Single-Cell cDNA Library Quantification and Quality Control

Single-Cell cDNA Library Quantification

1. Pipette 49 μL assay solution per well in black 96-well assay plates for samples. Keep the assay plates in the dark.
2. Collect the single-cell cDNA libraries by centrifugation of the storage plates for 15 sec at <2000 g. Carefully remove the sealing sheet.
3. Column by column, transfer 1 μL of each single-cell cDNA library directly into the assay solution in a black assay plate. Keep the assay plate in the dark.
4. After completing the transfer, seal the storage plate with the single-cell cDNA libraries. Collect them at the bottom of the well by centrifugation of the storage plates for 15 sec at <2000 g, and store at -20°C.
5. Prepare the standards for assay calibration. Place 47.5 μL assay solution per well and add 2.5 μL of the standards. We use four replicates of each standard 1 (0 ng/μL dsDNA, background) and standard 2 (10 ng/μL dsDNA, upper boundary).
6. After preparing both samples and the standards, we measure the fluorescence signal in a spectrofluorometer (excitation at 485 nm/emission at 530 nm). The concentration of the single-cell cDNA library is calculated according to the following equation with the dilution factor f_Dilution=50 and the average values of the standards.
7. After completion of measurements, discard the assay plates.

cDNA Quality Control

1. For each single-cell cDNA pool, prepare an eight-tube strip.
2. Collect the single-cell cDNA libraries by centrifugation of the storage plates for 15 sec at <2000 g. Carefully remove the sealing sheet.
3. Column by column, combine 1 μL of each single-cell cDNA library into an eight-tube strip.
4. After completing the transfer, seal the storage plate with the single-cell cDNA libraries. Collect them at the bottom of the well by centrifugation of the storage plates for 15 sec at <2000 g, and store at -20°C.
5. Combine the contents from each tube of the eight-tube strip into a single tube. This is the pooled single-cell cDNA library sample for profiling on the Bioanalyzer.
6. Follow the High Sensitivity DNA kit instructions (or similar), and analyze the cDNA profile traces.

Reference:

1. Schweingruber C, Nijssen J, Benitez J A, et al. Single-Cell mRNA-Seq of In Vitro-Derived Human Neurons Using Smart-Seq2[M]//Transcription Factor Regulatory Networks. Humana, New York, NY, 2023: 143-164.