Cell Biology

Salmonella enterica serovar Typhimurium (S. Typhimurium) is a Gram-negative, facultative intracellular bacterium, which causes gastrointestinal disorders in humans, and systemic, typhoid fever-like infections in mice. Our current knowledge regarding the involvement of cellular and humoral immunity in the defense from S. Typhimurium infections is largely based on animal models with attenuated strains. Cells of the innate immune system act as one of the first barriers in the defense from bacteria. We established a robust experimental model for the characterization of these cell types and their response during host-pathogen interactions. Therefore, this protocol focuses on the characterization of macrophages, monocytes, and neutrophils in the spleens of infected animals by employing multi-color flow cytometry.

Spontaneous DNA damage frequently occurs on the human genome, and it could alter gene expression by inducing mutagenesis or epigenetic changes. Therefore, it is highly desired to profile DNA damage distribution on the human genome and identify the genes that are prone to DNA damage. Here, we present a novel single-cell whole-genome amplification method which employs linear-copying followed by a split-amplification scheme, to efficiently remove amplification errors and achieve accurate detection of DNA damage in individual cells. In comparison to previous methods that measure DNA damage, our method uses a next-generation sequencing platform to detect misincorporated bases derived from spontaneous DNA damage with single-cell resolution.

In vivo erythropoiesis occurs in the erythroblast island niche (EBI), comprising of a central macrophage that attaches to and aids the maturation of erythroid progenitors into mature reticulocytes. Macrophages in hematopoietic tissue such as embryonic fetal liver are heterogeneous and express the cell surface protein F4/80. Earlier methods of isolating F4/80+ macrophages from hematopoietic tissue relied on FACS sorting, but the relatively low numbers of F4/80+ cells obtained after FACS sometimes led to poor RNA quality. Additionally, since EBI macrophages are attached to erythroblasts, care must be taken to avoid contamination with bound erythroblasts. We have developed a novel method for isolating F4/80+ cells from E13.5 mouse fetal liver using magnetic nanoparticles, which can be performed on the lab bench. During cell suspension and homogenization, we also add a peptide that disrupts erythroid macrophage interactions and generates F4/80+ single cells free of erythroid contamination. Thus, our protocol generates a population enriched in F4/80+ cells that are healthy and ready for sensitive techniques such as single cell sequencing.

The technology of cell carriers was developed as a response to the need for high cell density to enable higher production levels in cell-based production processes. To follow the production process, quantifying the number of cells on these carriers is required, as well as tracking their viability and proliferation. However, owing to various carriers’ unique structures, tracking the cells is challenging using current traditional assays that were originally developed for monolayers of adherent cells. The current "gold standard" method is counting cell nuclei, which is tedious and counts both live and dead cells. A few other techniques have been developed, but they are all specific to a carrier type and involve specialized equipment. Here, we describe a broad ranging method for counting cells on carriers. The method is based on the Alamar blue dye, a well-known, common marker for cell activity. No separation of the cells from the carriers is needed, nor is any specialized equipment. The method is simple and rapid, and provides comprehensive details necessary for control of production processes in cells. This method can be easily implemented in any cell-based process and other unique platforms for measuring growth of cells.

Graphic abstract:

Schematic of the in situ quantification method.

The formation of spheroids with mesenchymal stem/stromal cells (MSCs), mesenchymal bodies (MBs), is usually performed using bioreactors or conventional well plates. While these methods promote the formation of a large number of spheroids, they provide limited control over their structure or over the regulation of their environment. It has therefore been hard to elucidate the mechanisms orchestrating the structural organization and the induction of the trophic functions of MBs until now. We have recently demonstrated an integrated droplet-based microfluidic platform for the high-density formation and culture of MBs, as well as for the quantitative characterization of the structural and functional organization of cells within them. The protocol starts with a suspension of a few hundred MSCs encapsulated within microfluidic droplets held in capillary traps. After droplet immobilization, MSCs start clustering and form densely packed spherical aggregates that display a tight size distribution. Quantitative imaging is used to provide a robust demonstration that human MSCs self-organize in a hierarchical manner, by taking advantage of the good fit between the microfluidic chip and conventional microscopy techniques. Moreover, the structural organization within the MBs is found to correlate with the induction of osteo-endocrine functions (i.e., COX-2 and VEGF-A expression). Therefore, the present platform provides a unique method to link the structural organization in MBs to their functional properties.

Graphic abstract:

Droplet microfluidic platform for integrated formation, culture, and characterization of mesenchymal bodies (MBs). The device is equipped with a droplet production area (flow focusing) and a culture chamber that enables the culture of 270 MBs in parallel. A layer-by-layer analysis revealed a hierarchical developmental organization within MBs.

Experimental results in fungal biology research are usually obtained as average measurements across whole populations of cells, whilst ignoring what is happening at the single cell level. Microscopy has allowed us to study single-cell behavior, but it has low throughput and cannot be used to select individual cells for downstream experiments. Here we present a method that allows for the analysis and selection of single fungal cells in high throughput by flow cytometry and fluorescence activated cell sorting (FACS), respectively. This protocol can be adapted for every fungal species that produces cells of up to 70 microns in diameter. After initial setting of the flow cytometry gates, which takes a single day, accurate single cell analysis and sorting can be performed. This method yields a throughput of thousands of cells per second. Selected cells can be subjected to downstream experiments to study single-cell behavior.

Mammalian cell transfection is a powerful technique commonly used in molecular biology to express exogenous DNA or RNA in cells and study gene and protein function. Although several transfection strategies have been developed, there is a wide variation with regards to transfection efficiency, cell toxicity and reproducibility. Thus, a sensitive and robust method that can optimize transfection efficiency based not only on expression of the target protein of interest but also on the uptake of the nucleic acids, can be an important tool in molecular biology. Herein, we present a simple, rapid and robust flow cytometric method that can be used as a tool to optimize transfection efficiency while overcoming limitations of prior established methods that quantify transfection efficiency.

The ability to conduct investigation of cellular transcription, signaling, and function at the single-cell level has opened opportunities to examine heterogeneous populations at unprecedented resolutions. Although methods have been developed to evaluate high-dimensional transcriptomic and proteomic data (relating to cellular mRNA and protein), there has not been a method to evaluate corresponding high-dimensional functionomic data (relating to cellular functions) from single cells. Here, we present a protocol to quantitatively measure the differentiation potentials of single human hematopoietic stem and progenitor cells, and then cluster the cells according to these measurements. High dimensional functionomic analysis of cell potential allows cell function to be linked to molecular mechanisms within the same progenitor population.

Seedless vascular plants, including ferns and lycophytes, produce spores to initiate the gametophyte stage and to complete sexual reproduction. Approximately 10% of them are apomictic through the production of genomic unreduced spores. Being able to measure the spore nuclear DNA content is therefore important to infer their reproduction mode. Here we present a protocol of spore flow cytometry that allows an efficient determination of the reproductive modes of seedless vascular plants.

The main obstacle to eradicating HIV-1 from patients is post-integration latency (Finzi et al., 1999). Antiretroviral treatments target only actively replicating virus, while latent infections that have low or no transcriptional activity remain untreated (Sedaghat et al., 2007). To eliminate viral reservoirs, one strategy focuses on reversing HIV-1 latency via ‘shock and kill’ (Deeks, 2012). The basis of this strategy is to overcome the molecular mechanisms of HIV-1 latency by therapeutically inducing viral gene and protein expression under antiretroviral therapy and to cause selective cell death via the lytic properties of the virus, or the immune system now recognizing the infected cells. Recently, a number of studies have described the therapeutic potential of pharmacologically inhibiting members of the bromodomain and extraterminal (BET) family of human bromodomain proteins (Filippakopoulos et al., 2010; Dawson et al., 2011; Delmore et al., 2011) that include BRD2, BRB3, BRD4 and BRDT. Small-molecule BET inhibitors, such as JQ1 (Filippakopoulos et al., 2010; Delmore et al., 2011), I-BET (Nicodeme et al., 2010), I-Bet151 (Dawson et al., 2011), and MS417 (Zhang et al., 2012) successfully activate HIV transcription and reverse viral latency in clonal cell lines and certain primary T-cell models of latency. To identify the mechanism by which BET proteins regulate HIV-1 latency, we utilized small hairpin RNAs (shRNAs) that target BRD2, BRD4 and Cyclin T1, which is a component of the critical HIV-1 cofactor positive transcription elongation factor b (P-TEFb) and interacts with BRD2, and tested them in the CD4⁺ J-Lat A2 and A72 cell lines. The following protocol describes a flow cytometry-based method to determine the amount of transcriptional activation of the HIV-1 LTR upon shRNA knockdown. This protocol is optimized for studying latently HIV-1-infected Jurkat (J-Lat) cell lines.