Single cell RNA sequencing approaches are instrumental in studies of cell-to-cell variability. 5′ selective transcriptome profiling approaches allow simultaneous definition of the transcription start size and have advantages over 3′ selective approaches which just provide internal sequences close to the 3′ end. The only currently existing 5′ selective approach requires costly and labor intensive fragmentation and cell barcoding after cDNA amplification. Researchers at the CNRS Institute of Molecular & Cellular Pharmacology developed an optimized 5′ selective workflow where all the cell indexing is done prior to fragmentation. With their protocol, cell indexing can be performed in the Fluidigm C1 microfluidic device, resulting in a significant reduction of cost and labor. The researchers also designed optimized unique molecular identifiers that show less sequence bias and vulnerability towards sequencing errors resulting in an improved accuracy of molecule counting. They provide comprehensive experimental workflows for Illumina and Ion Proton sequencers that allow single cell sequencing in a cost range comparable to qPCR assays.
After cell lysis in 4.5 nl poly-adenylated RNA is reverse-transcribed in 31.5 nl with an anchored oligodT primer. A PCR primer sequence and unique molecular identifiers (UMIs) are added to the 3′ end of the cDNA via reverse transcriptase template switching. The cDNA is subsequently amplified and cell index sequences (barcode) as well as terminal biotins are introduced by PCR in the microfluidic device. The barcoded cDNAs are pooled, fragmented by tagmentation with Tn5 transposase and the biotinylated terminal fragments are isolated on streptavidin beads. 5′ terminal fragments are selectively amplified and additional sequences required for Ion Torrent sequencers are introduced by PCR.