A long-sought milestone in microfluidics research has been the development of integrated technology for scalable analysis of transcription in single cells. Here we present a fully integrated microfluidic device capable of performing high-precision RT-qPCR measurements of gene expression from hundreds of single cells per run. Our device executes all steps of single-cell processing, including cell capture, cell lysis, reverse transcription, and quantitative PCR. In addition to higher throughput and reduced cost, we show that nanoliter volume processing reduced measurement noise, increased sensitivity, and provided single nucleotide specificity. We apply this technology to 3,300 single-cell measurements of ( i ) miRNA expression in K562 cells, ( ii ) coregulation of a miRNA and one of its target transcripts during differentiation in embryonic stem cells, and ( iii ) single nucleotide variant detection in primary lobular breast cancer cells. The core functionality established here provides the foundation from which a variety of on-chip single-cell transcription analyses will be developed.

Single-cell genomics is critical for understanding cellular heterogeneity in cancer, but existing library preparation methods are expensive, require sample preamplification and introduce coverage bias. Here we describe direct library preparation (DLP), a robust, scalable, and high-fidelity method that uses nanoliter-volume transposition reactions for single-cell whole-genome library preparation without preamplification. We examined 782 cells from cell lines and triple-negative breast xenograft tumors. Low-depth sequencing, compared with existing methods, revealed greater coverage uniformity and more reliable detection of copy-number alterations. Using phylogenetic analysis, we found minor xenograft subpopulations that were undetectable by bulk sequencing, as well as dynamic clonal expansion and diversification between passages. Merging single-cell genomes in silico, we generated 'bulk-equivalent' genomes with high depth and uniform coverage. Thus, low-depth sequencing of DLP libraries may provide an attractive replacement for conventional bulk sequencing methods, permitting analysis of copy number at the cell level and of other genomic variants at the population level.

Heterogeneity in cell populations poses a major obstacle to understanding complex biological processes. Here we present a microfluidic platform containing thousands of nanoliter-scale chambers suitable for live-cell imaging studies of clonal cultures of nonadherent cells with precise control of the conditions, capabilities for in situ immunostaining and recovery of viable cells. We show that this platform mimics conventional cultures in reproducing the responses of various types of primitive mouse hematopoietic cells with retention of their functional properties, as demonstrated by subsequent in vitro and in vivo (transplantation) assays of recovered cells. The automated medium exchange of this system made it possible to define when Steel factor stimulation is first required by adult hematopoietic stem cells in vitro as the point of exit from quiescence. This technology will offer many new avenues to interrogate otherwise inaccessible mechanisms governing mammalian cell growth and fate decisions.

We present a microfluidic 'megapixel' digital PCR device that uses surface tension-based sample partitioning and dehydration control to enable high-fidelity single DNA molecule amplification in 1,000,000 reactors of picoliter volume with densities up to 440,000 reactors cm(-2). This device achieves a dynamic range of 10(7), single-nucleotide-variant detection below one copy per 100,000 wild-type sequences and the discrimination of a 1% difference in chromosome copy number.

Here we present an integrated microfluidic device for the high-throughput digital polymerase chain reaction (dPCR) analysis of single cells. This device allows for the parallel processing of single cells and executes all steps of analysis, including cell capture, washing, lysis, reverse transcription, and dPCR analysis. The cDNA from each single cell is distributed into a dedicated dPCR array consisting of 1020 chambers, each having a volume of 25 pL, using surface-tension-based sample partitioning. The high density of this dPCR format (118,900 chambers/cm(2)) allows the analysis of 200 single cells per run, for a total of 204,000 PCR reactions using a device footprint of 10 cm(2). Experiments using RNA dilutions show this device achieves shot-noise-limited performance in quantifying single molecules, with a dynamic range of 10(4). We performed over 1200 single-cell measurements, demonstrating the use of this platform in the absolute quantification of both high- and low-abundance mRNA transcripts, as well as micro-RNAs that are not easily measured using alternative hybridization methods. We further apply the specificity and sensitivity of single-cell dPCR to performing measurements of RNA editing events in single cells. High-throughput dPCR provides a new tool in the arsenal of single-cell analysis methods, with a unique combination of speed, precision, sensitivity, and specificity. We anticipate this approach will enable new studies where high-performance single-cell measurements are essential, including the analysis of transcriptional noise, allelic imbalance, and RNA processing.

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Recent advances in single-cell molecular analytical methods and clonal growth assays are enabling more refined models of human hematopoietic lineage restriction processes to be conceptualized. Here, we report the results of integrating single-cell proteome measurements with clonally determined lymphoid, neutrophilic/monocytic, and/or erythroid progeny outputs from >1000 index-sorted CD34+ human cord blood cells in short-term cultures with and without stromal cells. Surface phenotypes of functionally examined cells were individually mapped onto a molecular landscape of the entire CD34+ compartment constructed from single-cell mass cytometric measurements of 14 cell surface markers, 20 signaling/cell cycle proteins, and 6 transcription factors in ∼300 000 cells. This analysis showed that conventionally defined subsets of CD34+ cord blood cells are heterogeneous in their functional properties, transcription factor content, and signaling activities. Importantly, this molecular heterogeneity was reduced but not eliminated in phenotypes that were found to display highly restricted lineage outputs. Integration of the complete proteomic and functional data sets obtained revealed a continuous probabilistic topology of change that includes a multiplicity of lineage restriction trajectories. Each of these reflects progressive but variable changes in the levels of specific signaling intermediates and transcription factors but shared features of decreasing quiescence. Taken together, our results suggest a model in which increasingly narrowed hematopoietic output capabilities in neonatal CD34+ cord blood cells are determined by a history of external stimulation in combination with innately programmed cell state changes.

SummaryThe role of growth factors (GFs) in controlling the biology of human hematopoietic stem cells (HSCs) remains limited by a lack of information concerning the individual and combined effects of GFs directly on the survival, Mitogenesis, and regenerative activity of highly purified human HSCs. We show that the initial input HSC activity of such a purified starting population of human cord blood cells can be fully maintained over a 21-day period in serum-free medium containing five GFs alone. HSC survival was partially supported by any one of these GFs, but none were essential, and different combinations of GFs variably stimulated HSC proliferation. However, serial transplantability was not detectably compromised by many conditions that reduced human HSC proliferation and/or survival. These results demonstrate the dissociated control of these three human HSC bio-responses, and set the stage for future improvements in strategies to modify and expand human HSCs ex vivo.