Stem Cell

The development of human organotypic models of cartilage provides essential insights into chondrogenesis and chondrocyte hypertrophy while enabling advanced applications in drug discovery, gene editing, and tissue regeneration. Here, we present a robust and efficient protocol for differentiating human expanded pluripotent stem cells (hEPSCs) into hypertrophic chondrocytes through a sclerotome intermediate. The protocol involves initial sclerotome induction, followed by 3D chondrogenic culture and subsequent hypertrophic maturation induced by bone morphogenetic protein-4 (BMP4), thyroid hormone (T3), and β-glycerophosphate. This protocol also allows for sensitive testing of the effects of various compounds on hypertrophic differentiation during the maturation process. Furthermore, we identify an α-adrenergic receptor antagonist, phentolamine, as an inhibitor of hypertrophic differentiation. This organoid system provides a practical platform for exploring cartilage hypertrophy mechanisms and testing therapeutic strategies for cartilage regeneration.

Mouse embryonic fibroblasts (MEFs) derived from genetically modified mice are a valuable resource for studying gene function and regulation. The MEF system can also be combined with rescue studies to characterize the function of mutant genes/proteins, such as disease-causing variants. However, primary MEFs undergo senescence soon after isolation and passaging, making long-term genetic manipulations difficult. Previously described methods for MEF immortalization are often inconsistent or alter the physiological properties of the cells. Here, we describe an optimized method that overcomes these limitations. By using electroporation to deliver CRISPR constructs that target the Tp53 gene, the method reliably generates immortalized MEFs (iMEFs) within three weeks. Importantly, iMEFs closely resemble the parent cell populations, and individual iMEFs can be cloned and expanded for subsequent genetic manipulation and characterization. We envision that this protocol can be adopted broadly to immortalize other mouse primary cell types.

The blastocysts consist of dozens of cells of three distinct lineages: epiblast (Epi), trophoblast (TB), and primitive endoderm (PrE). All embryonic and extraembryonic tissues are derived from Epi, TB, and PrE. Stem cell lines representing preimplantation Epi and TB have been established and are known as embryonic stem cells (ESCs) and trophoblast stem cells (TSCs). Extraembryonic endoderm cells (XENCs) constitute a cell line that has been established from PrE. Although in vivo, PrE gives rise to visceral endoderm (VE), parietal endoderm (PE), and marginal zone endoderm (MZE); XENCs, on blastocyst injection into chimeras, primarily contribute to the distal region of PE. Here, we provide a comprehensive protocol for the establishment of fully potent primitive endoderm stem cell (PrESC) lines. PrESCs are established and maintained on mouse embryonic fibroblast (MEF) feeder cells in a serum-free medium supplemented with fibroblast growth factor 4 (FGF4), heparin, CHIR99021, and platelet-derived growth factor-AA (PDGF-AA). PrESCs co-express markers indicative of pluripotency and endoderm lineage commitment, exhibiting characteristics akin to those of PrE. On transplantation of PrESCs into blastocysts, they demonstrate a high efficiency in contributing to VE, PE, and MZE. PrESCs serve as a valuable model for studying PrE, sharing similarities in gene expression profiles and differentiation potential. PrESCs constitute a pivotal cornerstone for in vitro analysis of early developmental mechanisms and for studies of embryo reconstitution in vitro, particularly in conjunction with ESCs and TSCs.

Key features

• Establishment and maintenance of primitive endoderm stem cell (PrESCs) capable of recapitulating the developmental prowess inherent to PrE.

• Offering a source of PrE lineage for embryo-like organoid reconstitution studies.

Graphical overview

Embryonic development is a complex process integrating cell fate decisions and morphogenesis in a spatiotemporally controlled manner. Previous studies with model organisms laid the foundation of our knowledge on post-implantation development; however, studying mammalian embryos at this stage is a difficult and laborious process. Early attempts to recapitulate mammalian development in vitro begun with embryoid bodies (EBs), in which aggregates of mouse embryonic stem cells (mESCs) were shown to differentiate into spatially arranged germ layers. A more revised version of EBs, gastruloids, improved the germ layer differentiation efficiency and demonstrated cell fate patterning on multiple axes. However, gastruloids lack anterior neural progenitors that give rise to brain tissues in the embryo. Here, we report a novel culture protocol to coax mESCs into post-implantation epiblast-like (EPI) aggregates in high throughput on bioengineered microwell arrays. We show that upon inhibition of the Wnt signaling pathway, EPI aggregates establish an extended axial patterning, leading to co-derivation of anterior neural progenitors and posterior tissues. Our approach is amenable to large-scale studies aimed at identifying novel regulators of gastrulation and anterior neural development that is currently out of reach with existing embryoid models. This work should contribute to the advancement of the nascent field of synthetic embryology, opening up exciting perspectives for various applications of pluripotent stem cells in disease modeling and tissue engineering.

Key features

• A new gastruloid culture system to model post-implantation mouse embryonic development in vitro

• High-throughput formation of epiblast-like aggregates on hydrogel microwells

• Builds upon conventional gastruloid cultures and provides insight into the role of Wnt signaling for the formation of anterior neural tissues

Graphical overview

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.

Graphical abstract:

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.

Graphical abstract:

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.

Graphical abstract:

Macrophages are key cells in the innate immune system and play a role in a variety of diseases. However, macrophages are terminally differentiated and difficult to manipulate genetically via transfection or through CRISPR-Cas9 gene editing. To overcome this limitation, we provide a simplified protocol for the generation of mouse embryonic stem cells-derived macrophages (ESDM). Thus, genetic manipulation can be performed using embryonic stem cells, selecting for the desired changes, and finally producing macrophages to study the effects of the previous genetic manipulation. These studies can contribute to many areas of research, including atherosclerosis and inflammation. Production of ESDM has been previously achieved using embryoid body (EB) intermediates. Here, we optimized the EB method using a simplified medium, reducing the number of recombinant proteins and medium recipes required. Our EB-based differentiation protocol consists of three stages: 1) floating EB formation; 2) adherence of EBs and release of floating macrophage progenitors; and, 3) terminal differentiation of harvested macrophage progenitors. The advantages of this protocol include achieving independent floating EBs in stage 1 by using a rocker within the tissue culture incubator, as well as the exclusion of small EBs and cell clusters when harvesting macrophage progenitors via cell filtration.

Extracellular vesicles (EVs) are thought to mediate intercellular communication through the delivery of cargo proteins and RNA to target cells. The uptake of EVs is often followed visually using lipophilic-dyes or fluorescently-tagged proteins to label membrane constituents that are then internalized into recipient cells (Christianson et al., 2013; De Jong et al., 2019). However, these methods do not probe the exposure of EV cargo to intracellular compartments, such as the cytoplasm and nucleus, where protein or RNA molecules could elicit functional changes in recipient cells. In this protocol, we employ an EV cargo protein-APEX fusion to detect proximity interactions with recipient cell cytoplasmic/nuclear targets. This approach results in the biotinylation of proteins in close contact with the reporter fusion and thus permits profiling of biotinylated proteins affinity purified on immobilized streptavidin beads.

Graphic abstract:

Schematic showing three steps of APEX-mediated proximity labeling of proteins in cells targeted by EVs.

Inducing loss of function of a target protein using methods such as gene knockout is a powerful and useful strategy for analyzing protein function in cells. In recent years, the CRISPR/Cas-9-based gene knockout technology has been widely used across a variety of eukaryotes; however, this type of simple gene knockout strategy is not applicable to essential genes, which require a conditional knockout system. The auxin-inducible degron (AID) system enables rapid depletion of the target protein in an auxin-dependent manner and has been used to generate conditional mutants in various eukaryotic cell lines. One problem with the AID system is the use of high auxin concentrations for protein degradation, which can cause cytotoxicity. Recently, we established a super-sensitive AID (ssAID) system that allowed a reduction in the amount of auxin required by more than 1,000-fold. We also utilized a single-step method to generate AID-based conditional knockout cells with a ssAID system in various cell lines. In this protocol, we introduce our improved method, which provides a powerful tool for the investigation of the roles of essential genes.