FLASH-seq Low-Amplification protocol

Prepare the following lysis mix:

A	B	C	D
Reagent	Reaction concentration	Volume (µl)	384-well plate
Triton-X100 (10% v/v)	0.2%	0.020	8.448
dNTP mix (25 mM each)	6 mM	0.240	101.376
SMART dT30VN (100 µM)	1.8 µM	0.018	7.603
RNAse inhibitor (40 U/µl)	1.2 U/µl	0.030	12.672
DTT (100 mM)	1.2 mM	0.012	5.069
FS TSO (100 µM)	9.2 µM	0.092	38.861
dCTP (100 mM)	9 mM	0.090	38.016
Betaine (5 M)	1 M	0.200	84.480
Nuclease-free water	-	0.298	125.875
Total volume (µl)		1.000	422.400

Add 1µL lysis mix to each well of a 384-well plate

Seal the plate with a PCR seal and quickly spin it down to collect the lysis mix to the bottom.

Proceed immediately to the next step or store the plate at -20°C long-term. Plates that are going to be used on the same day can be stored in the fridge or kept on ice.

Sort single cells into 384-well plates containing 1µL lysis mix.

Seal the plate with an aluminium seal. If processing multiple plates at once, keep each plate on dry ice until ready to transfer them all at -80°C for long-term storage. Plates containing single cells should ideally be processed within 6 months.

Remove the plates from the -80°C freezer and check that the aluminium seal is still intact. If damaged or not sticking to the plate anymore, wait a few minutes for the plate to partially thaw, remove the damaged foil and replace it with a new one.

Place the plate in a thermocycler with a heated lid and incubate for 0h 3m 0s at 72°C , followed by a 4°C hold step.

Spin down any condensation droplets that may have formed during the incubation and return the plate to a cool rack. Proceed quickly to the next step. If not ready with the RT-PCR mix, keep the plate on the cool rack at all times.

While the plate is in the thermocycler, prepare the following RT-PCR mix:

A	B	C	D
Reagent	Reaction concentration	Volume (µl)	384-well plate
DTT (0.1 M)	4.8 mM	0.238	100.531
MgCl2 (1 M)	9.2 mM	0.046	19.430
Betaine (5 M)	800 mM	0.800	337.920
RNAse inhibitor (40 U/µl)	0.8 U/µl	0.096	40.550
SuperScript IV (200 U/µl)	2.00 U/µl	0.050	21.120
KAPA HiFi HotStart ReadyMix (2 x)	1 x	2.500	1056.000
Nuclease-free water	-	0.270	114.048
Total volume (µl)		4.000	1689.600

Add 4µL RT-PCR mix into each well of the 384-well plate.

Seal the plate with a PCR seal, gently vortex and spin down to collect the liquid at the bottom.

Place it in a thermocycler with heated lid and start the following RT-PCR program:

A	B	C	D	E
Step		Temperature	Time	Cycles
RT		50ºC	60 min	1 x
PCR	initial denaturation	98ºC	3 min	1 x
	denaturation	98ºC	20 sec	10-16 x*
	annealing	67ºC
	elongation	72ºC
		15ºC	Hold

*Adjust the number of cycles according to the cell type. We recommend 10-12 cycles for HEK 293T cells and 14-16 cycles for hPBMC.

Please note that the Tn5 transposase amount is a suggested starting point only. Optimisation might be necessary, depending on the specific activity of each batch of Tn5 and desired library size.

Indexing primers can be purchased from Illumina (Nextera XT index kit v2) or ordered from your local oligo manufacturer. In the "Materials" section we have added additional sequences for higher multiplexing.

Prepare the tagmentation mix as described below:

A	B	C
Reagent	Volume (µl)	Final concentration
TAPS-Mg buffer, pH=7.3 (5x)	2.000	10 mM TAPS, 5 mM MgCl2
Dimethylformamide (DMF) (100%)	2.000	20%
Tn5 transposase (2 µM working dil.)	0.025	5 nM
Nuclease-free water	4.975
Total volume (µl)	9.000

Dispense 9µL tagmentation mix in a new 384-well plate.

Add 1µL unpurified cDNA to each well containing the tagmentation mix.

Seal the plate, vortex, spin down, and carry out the tagmentation reaction: 55°C for 0h 8m 0s , 4°C hold. Upon completion proceed immediately to the next step.

Add 2.5µL 0.2% SDS to each well. Seal the plate, vortex, spin down and incubate 5 min at room temperature. Do not put the plate back on ice.

Add 2.5µL N7xx + S5xx index adaptors (5micromolar (µM) each).

Add 10µL enrichment PCR mix to each well:

A	B	C
Reagent	Volume (µl)	Final concentration
KAPA HiFi enzyme (1 U/μl)	0.50	0.02 U/μl
KAPA HiFi Buffer (5 x)	5.00	1 x
dNTPs (10 mM)	0.75	300 nM
Nuclease-free water	3.75
Total volume (µl)	10.00

Seal the plate, vortex, spin down, and place it in a thermocycler and carry out the enrichment PCR reaction. Adjust the number of PCR cycles according to the number of processed cells AND the number of pre-amplification cycles used in the RT-PCR reaction.

A	B	C	D	E
Step		Temperature	Time	Cycles
gap filling		72ºC	3 min	1 x
enrichment PCR	initial denaturation	98ºC	30 sec	1 x
	denaturation	98ºC	10 sec	14-16 x
	annealing	55ºC	30 sec
	elongation	72ºC	30 sec
		15ºC	hold

Take an aliquot from each sample for the final library cleanup (i.e. 5 µl). and transfer it to a 1.5-ml Eppendorf tube. The rest of the library can be stored long-term at -20°C .

Remove the Sera-Mag SpeedBeads™ working solution from the 4°C storage and equilibrate it at room temperature for 0h 15m 0s .

Add Sera-Mag SpeedBeads™ working solution to a final ratio of 0.8 x and mix well to homogenisation.

Incubate the tube off the magnetic stand for 0h 5m 0s at Room temperature .

Place the tube on the magnetic stand and leave it for 0h 5m 0s or until the solution appears clear.

Remove the supernatant without disturbing the beads.

Recommended: wash the pellet with 1mL 80% v/v ethanol. Incubate 0h 0m 30s without removing the tube from the magnetic stand.

Remove any trace of ethanol and let the bead pellet dry for 0h 2m 0s or until small cracks appear. Do not cap the tube or remove it from the magnetic stand during this time. Do not completely air-dry the beads.

Remove the tube from the magnetic stand, add 50µL nuclease-free water and mix well by pipetting or vortexing to resuspend the beads.

Incubate 0h 2m 0s off the magnetic stand.

Place the tube back on the magnetic stand and incubate for 0h 2m 0s or until the solution appears clear.

Remove 49µL of the supernatant and transfer it to a new 1.5-ml LoBind tube. Store the cDNA at -20°C long-term or until ready for sequencing.

Use Qubit fluorometer to quantify the library. Library yield can vary depending on the number of cells being pooled.

Check the final library size on the Agilent Bioanalyzer.

Use the average size indicated on the Bioanalyzer and the concentration reported after Qubit measurement to determine the exact molarity required for sequencing.

The purified library can be sequenced on any Illumina sequencer. Follow the specifications reported for each instrument. Single-end 75 bp is generally sufficient but longer read modes or paired-end sequencing can be an option, depending on the question at hand.

These instructions briefly describe the data processing of the sequencing results. The final pipeline will likely have to be adapted to the question at hand. The following lines assume that all the programs and their dependencies are installed on your machine and that the data are single-end reads (75 bp). Some values, such as the number of threads and RAM usage may have to be adapted to your machine settings.

It should be noted that there are many other ways to analyse full-length single-cell RNA-sequencing data. Pseudo-alignment tools (e.g., Salmon or Kallisto) or automatic pipelines (zUMIs) could be used as well.

Requirements (tested version):

• bcl2fastq (v2.20)
• STAR (v2.7.3)
• FeatureCounts (v1.6.5)
• BBMAP (v38.86)
• samtools (v1.9)
• IGV

Sample demultiplexing

Sequencing results will be delivered as demultiplexed FASTQ or raw bcl2 files. To convert bcl2 files to FASTQ, bcl2fastq program (Illumina) can be used.

# 0. Variables
BASECALL_DIR="/path/to/flowcell/Data/Intensities/BaseCalls/"
OUTPUT_DIR="/path/to/output_folder/"
SAMPLESHEET="/path/to/Demultiplexing_SampleSheet.csv"

# 1. Bcl2fastq
ulimit -n 10000
cd /path/to/flowcell/
bcl2fastq --input-dir $BASECALL_DIR --output-dir $OUTPUT_DIR --sample-sheet $SAMPLESHEET --create-fastq-for-index-reads --no-lane-splitting
```When sequencing on a NextSeq 550 instrument, the sample sheet should contain the following information in a csv file:



<img src="https://static.yanyin.tech/literature_test/protocol_io_true/protocols.io.yxmvmnod5g3p/Screenshot%202021-12-12%20at%2008.58.20.png" alt="" loading="lazy" title=""/>

Illumina Experiment Manager can be used to assist you in creating the sample sheet. 



We recommend exploring the barcode combinations left in the undetermined reads looking to confirm that all the cells have been properly demultiplexed.

zcat Undetermined_S0_I1_001.fastq.gz | awk -F' 1:N:0:' 'NR%4==1{print $2}' | sort | uniq -c > left_index.txt sort -k1,1 left_index.txt

for file in ./out/R1 do zcat $file | wc -l done

Index the genome

The reference genome needs to be indexed prior to any mapping. The FASTA and GTF references can be obtained from ENSEMBL, Gencode, UCSC, ...

# 0. Variables
OUTPUTREF="/path/to/STAR_indexed_genome/"
FASTA="GRCh38.primary_assembly.genome.fa"
GTF="gencode.v34.primary_assembly.annotation.gtf"

# 1. Genome indexing
# sjdbOverhang should be adapted based on the read length (read_length - 1)
mkdir $OUTPUTREF

STAR --runThreadN 15 --runMode genomeGenerate --genomeDir $OUTPUTREF --genomeFastaFiles $FASTA --sjdbGTFfile $GTF --sjdbOverhang 74

FASTQ trimming (optional)

If you observe sequencing primer left-overs the FASTQ files can be trimmed using BBDUK or Trimmomatic.

bbduk.sh -Xmx48g in=sample.fastq.gz out=cleaned.left.fastq t=32 ktrim=l ref=adapters.fa k=23 mink=7 hdist=1 hdist2=0 tbo
bbduk.sh -Xmx48g in=cleaned.left.fastq out=cleaned.fastq t=32 ktrim=r ref=adapters.fa k=23 mink=7 hdist=1 hdist2=0 tbo

mv FASTQ/cleaned.fastq FASTQ/sample.R1.fastq.gz

Mapping

The FASTQ file can then be mapped onto the reference genome. Example for one sample, use a loop or parallelise this task to process all the cells:

# 0. Variables
GENOME="/path/to/STAR_indexed_genome/"
FASTQ="/path/to/sample.R1.fastq.gz"
ID=”sample_id”

# 1. Mapping
STAR --runThreadN 30 --limitBAMsortRAM 20000000000 --genomeLoad LoadAndKeep --genomeDir "$GENOME" --readFilesIn "$FASTQ" --readFilesCommand zcat --limitSjdbInsertNsj 2000000 --outFilterIntronMotifs RemoveNoncanonicalUnannotated --outSAMtype BAM SortedByCoordinate --outFileNamePrefix "$ID"_

# 2. SAM to sorted BAM
# -F 260 filters out unmapped and secondary alignments
samtools view -@ 30 -Sb -F 260 "$ID"_Aligned.sortedByCoord.out.bam > "$ID"_Aligned.sortedByCoord.filtered.bam
samtools index "$ID"_Aligned.sortedByCoord.filtered.bam

Data visualization (optional)

Once the reads have been mapped we highly recommend using the Integrated Genome Viewer (IGV) to visualise the mapping results and ensure that the results make sense. As a quick check-up visualise a few housekeeping genes (i.e., ACTB, GAPDH, …) and cell specific markers to look for reads mapping to exon, intron, exon-intron junctions. Look for abnormalities such as read piles falling in intergenic or centromeric regions.

No single-cell RNA sequencing protocol is perfect and non-specific priming, genomic DNA contaminations, … can happen but should represent rare events.

Recurrent soft-clipping could also indicate the presence of sequencing adaptor left-overs that could affect the mapping rate.

Count matrix

Finally, t​he number of reads associated with each gene can be obtained as follows:

featureCounts -T 1 -t exon -g gene_name --fracOverlap 0.25 -a "$GTF" -o "$ID"_ReadCount.featureCounts.gencode.txt "$ID"_Aligned.sortedByCoord.filtered.bam

Post-processing

The post-processing steps will vary depending on the question at hand. The online book “Orchestrating Single-Cell Analysis with Bioconductor” (https://bioconductor.org/books/release/OSCA/) is a gold mine of information that can be used to help you design your own pipeline. Alternatively, Seurat (R, https://satijalab.org/seurat/) or scanpy (python, https://scanpy.readthedocs.io/en/stable/) provide tools compatible with FLASH-seq data. Given their similarities, we currently recommend using Smart-seq2 guidelines when processing FLASH-seq data.