Stem Cell

In mammals, the skin comprises several distinct cell populations that are organized into the following layers: epidermis (stratum corneum, stratum granulosum, stratum spinosum, and basal layer), basement membrane, dermis, and hypodermal (subcutaneous fat) layers. It is vital to identify the exact location and function of proteins in different skin layers. Laser capture microdissection (LCM) is an effective technique for obtaining pure cell populations from complex tissue sections for disease-specific genomic and proteomic analysis. In this study, we used LCM to isolate different skin layers, constructed a stratified developmental lineage proteome map of human skin that incorporates spatial protein distribution, and obtained new insights into the role of extracellular matrix (ECM) on stem cell regulation.

Primary hematopoietic stem and progenitor cell (HSPC)-derived megakaryocytes are a valuable tool for translational research interrogating disease pathogenesis and developing new therapeutic avenues for patients with hematologic disorders including myeloproliferative neoplasms (MPNs). Thrombopoietin (TPO)-independent proliferation and megakaryocyte differentiation play a central role in the pathogenesis of essential thrombocythemia and myelofibrosis, two MPN subtypes that are characterized by increased numbers of bone marrow megakaryocytes and somatic mutations in either JAK2, CALR, or MPL. However, current culture strategies generally use healthy HSPCs for megakaryocyte production and are not optimized for the investigation of TPO-independent or TPO-hypersensitive growth and megakaryocyte-directed differentiation of primary patient–derived HSPCs. Here, we describe a detailed protocol covering all necessary steps for the isolation of CD34⁺ HSPCs from the peripheral blood of MPN patients and the subsequent TPO-independent differentiation into CD41⁺ megakaryocytes using both a collagen-based colony assay and a liquid culture assay. This protocol provides a novel, reproducible, and cost-effective approach for investigating megakaryocyte growth and differentiation properties from primary MPN patient cells that can be easily adapted for research on other megakaryocyte-related disorders.

Graphical abstract

Schematic representation of the isolation of CD34⁺ progenitor cells and subsequent TPO-independent megakaryocyte differentiation

Sphingolipids are important structural components of cellular membranes. They also function as prominent signaling molecules to control a variety of cellular events, such as cell growth, differentiation, and apoptosis. Impaired sphingolipid metabolism, particularly defects in sphingolipid degradation, has been associated with many human diseases. Fluorescence sphingolipid analogs have been widely used as efficient probes to study sphingolipid metabolism and intracellular trafficking in living mammalian cells. Compared with nitrobenzoxadiazole fluorophores (NBD FL), the boron dipyrromethene difluoride fluorophores (BODIPY FL) have much higher absorptivity and fluorescence quantum. These features allow more intensive labeling of cells for fluorescence microscopy imaging and flow cytometry analysis. Here, we describe a protocol employing BODIPY FL-labeled sphingolipid analogs to elucidate sphingolipid internalization, trafficking, and endocytosis in mouse embryonic stem cells.

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Skeletal muscle stem cells differentiated from human-induced pluripotent stem cells (hiPSCs) serve as a uniquely promising model system for investigating human myogenesis and disease pathogenesis, and for the development of gene editing and regenerative stem cell therapies. Here, we present an effective and reproducible transgene-free protocol for derivation of human skeletal muscle stem cells, iMyoblasts, from hiPSCs. Our two-step protocol consists of 1) small molecule-based differentiation of hiPSCs into myocytes, and 2) stimulation of differentiated myocytes with growth factor-rich medium to activate the proliferation of undifferentiated reserve cells, for expansion and cell line establishment. iMyoblasts are PAX3⁺/MyoD1⁺ myogenic stem cells with dual potential to undergo muscle differentiation and to self-renew as a regenerative cell population for muscle regeneration both ex vivo and in vivo. The simplicity and robustness of iMyoblast generation and expansion have enabled their application to model the molecular pathogenesis of Facioscapulohumeral Muscular Dystrophy and Limb-Girdle Muscular Dystrophies, to both ex vivo and in vivo muscle xenografts, and to respond efficiently to gene editing, enabling the co-development of gene correction and stem cell regenerative therapeutic technologies for the treatment of muscular dystrophies and muscle injury.

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Limbal mesenchymal stromal cells (LMSC), a cellular component of the limbal stem cell niche, have the capability of determining the fate of limbal epithelial progenitor cells (LEPC), which are responsible for the homeostasis of corneal epithelium. However, the isolation of these LMSC has proven to be difficult due to the small fraction of LMSC in the total limbal population, and primary cultures are always hampered by contamination with other cell types. We recently published the efficient isolation and functional characterization of LMSC from the human corneal limbus using CD90 as a selective marker. We observed that flow sorting yielded a pure population of LMSC with superior self-renewal capacity and transdifferentiation potential, and supported the maintenance of the LEPC phenotype. Here, we describe an optimized protocol for the isolation of LMSC from cadaveric corneal limbal tissue by combined collagenase digestion and flow sorting with expansion of LMSC on plastic.

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To optimize differentiation protocols for stem cell-based in vitro modeling applications, it is essential to assess the change in gene expression during the differentiation process. This allows controlling its differentiation efficiency into the target cell types. While RNA transcriptomics provides detail at a larger scale, timing and cost are prohibitive to include such analyses in the optimization process. In contrast, expression analysis of individual genes is cumbersome and lengthy.

Here, we developed a versatile and cost-efficient SYBR Green array of 27 markers along with two housekeeping genes to quickly screen for differentiation efficiency of human induced pluripotent stem cells (iPSCs) into excitatory cortical neurons. We first identified relevant pluripotency, neuroprogenitor, and neuronal markers for the array by literature search, and designed primers with a product size of 80-120 bp length, an annealing temperature of 60°C, and minimal predicted secondary structures. We spotted combined forward and reverse primers on 96-well plates and dried them out overnight. These plates can be prepared in advance in batches and stored at room temperature until use. Next, we added the SYBR Green master mix and complementary DNA (cDNA) to the plate in triplicates, ran quantitative PCR (qPCR) on a Quantstudio 6 Flex, and analyzed results with QuantStudio software.

We compared the expression of genes for pluripotency, neuroprogenitor cells, cortical neurons, and synaptic markers in a 96-well format at four different time points during the cortical differentiation. We found a sharp reduction of pluripotency genes within the first three days of pre-differentiation and a steady increase of neuronal markers and synaptic markers over time. In summary, we built a gene expression array that is customizable, fast, medium-throughput, and cost-efficient, ideally suited for optimization of differentiation protocols for stem cell-based in vitro modeling.

Transplantation of hematopoietic material into recipient mice is an assay routinely used to determine the presence and function of hematopoietic stem and progenitor cells (HSPCs) in vivo. The principle of the method is to transplant donor cells being tested for HSPCs into a recipient mouse following bone marrow ablation and testing for reconstitution of hematopoiesis. Congenic mouse strains where donor and recipient differ by a distinct cell surface antigen (commonly CD45.1 versus CD45.2) are used to distinguish between cells derived from the donor and any residual recipient cells. Typically, the transplantation is performed using bone marrow cells, which are enriched for HSPCs. Here, we describe an analogous procedure using hematopoietic material from spleen, allowing detection of functional progenitors and/or stem cells in the spleen that can occur under certain pathologies. Key to the success of this procedure is the prior removal of mature T cells from the donor sample, to minimize graft versus host reactions. As such, this protocol is highly analogous to standard bone marrow transplant procedures, differing mainly only in the source of stem cells (spleen rather than bone marrow) and the recommendation for T cell depletion to avoid potential immune incompatibilities.

Graphical abstract:

Schematic overview for assessment of stem cells in spleen by transplantation.
Single cell suspensions from spleens are depleted of potentially pathogenic mature T lymphocytes by magnetic bead immunoselection using biotinylated antibodies against CD4 and CD8, followed by streptavidin magnetic beads, which are subsequently removed by using a magnet (MojoSort, Biolegend). Successful T cell depletion is then evaluated by Fluorescence Activated Cell Sorting (FACS). T-cell depleted cell suspension is injected intravenously through the retro-orbital sinus into lethally irradiated recipients. Recipients are analyzed for successful engraftment by FACS analysis for the presence of donor-derived mature hematopoietic lineages in the peripheral blood. A second serial transplantation can be used to document the presence of long-term reconstituting stem cells in the periphery of the original donor mice.

This protocol describes a method to assess relative changes in the level of global protein synthesis in the preimplantation embryo using the Click-iT^® Plus OPP Protein Synthesis Assays. In this assay, O-propargyl-puromycin (OPP), an analog of puromycin, is efficiently incorporated into the nascent polypeptide of newly translated proteins in embryonic cells. OPP is fluorescently labeled with a photostable Alexa Fluor^TM dye and detected with fluorescence microscopy. The intensity of the fluorescence is quantitatively analyzed. This is a fast, sensitive, and non-radioactive method for the detection of protein synthesis in early embryo development. It provides a tool for analyzing the temporal regulation of protein synthesis, as well as the effects of changes in the embryonic microenvironment, and pharmacological and genetic modulations of embryo development.

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Figure 1. Brief overview of the procedures of the Click-iT^® Plus OPP Alexa Fluor^® protein synthesis assay in embryonic cells.

(A) Set up OPP treatments: (1) Set up microdrops containing 50 µL of OPP working solution and label different treatments on the back of culture dishes (e.g., T0, T1, T2, and T3); (2) The drops are overlain with 2–3 mm heavy paraffin oil and then equilibrated in incubator for 2 h; (3) Collect the embryos from female reproductive tracts or following in vitro culture in desired treatments; (4) Culture embryos in the equilibrated OPP working solution for 2–6 h. (B) Example of OPP detection procedures working with 60-well plates labeled as T0, T1, T2, T3, T4, and T5 for different treatments: (1) The first 60-well plate is used for the procedures of washing, fixation, permeabilization, and Click-iT^® OPP detection. (2) The second 60-well plate is for DNA staining and washing. (C) Slide preparation: (1) Label the required number of slides and set up vaseline coverslip supports; (2) Add mounting medium; (3) Transfer embryos into mounting medium; (4) Set coverslip; (5) Seal the coverslip with nail polish.

Although CRISPR-Cas9 genome editing can be performed directly in single-cell mouse zygotes, the targeting efficiency for more complex modifications such as the insertion of two loxP sites, multiple mutations in cis, or the precise insertion or deletion of longer DNA sequences often remains low (Cohen, 2016). Thus, targeting and validation of correct genomic modification in murine embryonic stem cells (ESCs) with subsequent injection into early-stage mouse embryos may still be preferable, allowing for large-scale screening in vitro before transfer of thoroughly characterized and genetically defined ESC clones into the germline. This procedure can result in a reduction of animal numbers with cost effectiveness and compliance with the 3R principle of animal welfare regulations. Here, we demonstrate that after transfection of homology templates and PX458 CRISPR-Cas9 plasmids, EGFP-positive ESCs can be sorted with a flow cytometer for the enrichment of CRISPR-Cas9-expressing cells. Cell sorting obviates antibiotic selection and therefore allows for more gentle culture conditions and faster outgrowth of ESC clones, which are then screened by qPCR for correct genomic modifications. qPCR screening is more convenient and less time-consuming compared to analyzing PCR samples on agarose gels. Positive ESC clones are validated by PCR analysis and sequencing and can serve for injection into early-stage mouse embryos for the generation of chimeric mice with germline transmission. Therefore, we describe here a simple and straightforward protocol for CRISPR-Cas9-directed gene targeting in ESCs.

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Craniofacial anomalies (CFA) are a diverse group of deformities, which affect the growth of the head and face. Dysregulation of cranial neural crest cell (NCC) migration, proliferation, differentiation, and/or cell fate specification have been reported to contribute to CFA. Understanding of the mechanisms through which cranial NCCs contribute for craniofacial development may lead to identifying meaningful clinical targets for the prevention and treatment of CFA. Isolation and culture of cranial NCCs in vitro facilitates screening and analyses of molecular cellular mechanisms of cranial NCCs implicated in craniofacial development. Here, we present a method for the isolation and culture of cranial NCCs harvested from the first branchial arch at early embryonic stages. Morphology of isolated cranial NCCs was similar to O9-1 cells, a cell line for neural crest stem cells. Moreover, cranial NCCs isolated from a transgenic mouse line with enhanced bone morphogenetic protein (BMP) signaling in NCCs showed an increase in their chondrogenic differentiation capacity, suggesting maintenance of their in vivo differentiation potentials observed in vitro. Taken together, our established method is useful to visualize cellular behaviors of cranial NCCs.