Stem Cell

The development of patient-derived cardiac disease models has advanced rapidly due to the progress of human-induced pluripotent stem cell (hiPSC) technologies. Many protocols detail individual parts of the entire workflow, from handling hiPSCs and differentiating them into cardiomyocytes to live contraction imaging via widefield/phase-contrast and fluorescence microscopy. Here, we propose a streamlined protocol that guides users through hiPSC culture, differentiation, expansion, and functional imaging of hiPSC cardiomyocytes. First, hiPSC maintenance and handling procedures are outlined. Differentiation occurs over a two-week period, followed by selective expansion to increase the yield of hiPSC cardiomyocytes. Comprehensive characterization and quantification enable detailed contraction profiling of these cells. Designed to be low-cost, this protocol is suited for applications in drug discovery, screening, and clinical testing of patient-specific phenotypes. The addition of cardiomyocyte expansion and automated analysis distinguishes our protocol from current approaches.

Human induced pluripotent stem (iPS) cell lines harboring mutations in disease-related genes serve as invaluable in vitro models for unraveling disease mechanisms and accelerating drug discovery efforts. Introducing mutations into iPS cells using traditional gene editing approaches based on the CRISPR-Cas9 endonuclease often encounters challenges such as unintended insertions/deletions (indels) and off-target effects. To address these limitations, we present a streamlined protocol for introducing highly accurate gene mutations into human iPS cells using prime editing, a “search-and-replace” genome-editing technology that combines unwanted indel-minimized CRISPR-Cas9 nickase with reverse transcriptase. This protocol encompasses the design of prime editing guide RNAs (pegRNAs) required for binding and replacement at target loci, construction of prime editor and pegRNA expression vectors, gene transfer into iPS cells, and cell line selection. This protocol allows for the efficient establishment of disease-associated gene variants within 6–8 weeks while preserving critical genomic context.

Bone repair is a complex regenerative process relying on skeletal stem/progenitor cells (SSPCs) recruited predominantly from the periosteum. Activation and differentiation of periosteal SSPCs occur in a heterogeneous environment, raising the need for single cell/nucleus transcriptomics to decipher the response of the periosteum to injury. Enzymatic cell dissociation can induce a stress response affecting the transcriptome and lead to overrepresentation of certain cell types (i.e., immune and endothelial cells) and low coverage of other cell types of interest. To counteract these limitations, we optimized a protocol to isolate nuclei directly from the intact periosteum and from the fracture callus to perform single-nucleus RNA sequencing. This protocol is adapted for fresh murine periosteum, fracture callus, and frozen human periosteum. Nuclei are isolated using mechanical extraction combined with fluorescence-based nuclei sorting to obtain high-quality nucleus suspensions. This protocol allows the capture of the full diversity of cell types in the periosteum and fracture environment to better reflect the in vivo tissue composition.

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.

Recurrent hormone receptor-positive (HR+) breast cancer is a leading cause of cancer mortality in women. Recurrence and resistance to targeted therapies have been difficult to study due to the long clinical course of the disease, the complex nature of resistance, and the lack of clinically relevant model systems. Existing models are limited to a few HR+ cell lines, organoid models, and patient-derived xenograft models, all lacking components of the human tumor microenvironment. Furthermore, the low take rate and loss of estrogen receptor (ER) expression in patient-derived organoids (PDOs) has been challenging. Our protocol allows simultaneous isolation of PDOs and matching cancer-associated fibroblasts (CAFs) from primary and metastatic HR+ breast cancers. Importantly, our protocol has a higher take rate and enables long-term culturing of PDOs that retain ER expression. Our matching PDOs and CAFs will provide researchers with a new resource to study the influence of the tumor microenvironment on various aspects of cancer biology such as cell growth and drug resistance in HR+ breast cancer.

The human intestine plays a pivotal role in nutrient absorption and immune system regulation. Along the longitudinal axis, cell-type composition changes to meet the varying functional requirements. Therefore, our protocol focuses on the processing of the whole human intestine to facilitate the analysis of region-specific characteristics such as tissue architecture and changes in cell populations. We describe how to generate a biobank that can be used to isolate specific immune cell subtypes, generate organoid lines, and establish autologous immune cell-organoid co-cultures.

The advent of clustered regularly interspaced short palindromic repeats (CRISPR)/Cas9-based genome editing has marked a significant advancement in genetic engineering technology. However, the editing of induced pluripotent stem cells (iPSCs) with CRISPR presents notable challenges in ensuring cell survival and achieving high editing efficiency. These challenges become even more complex when considering the specific target site. P53 activation as a result of traditional CRISPR editing can lead to apoptosis, potentially worsening cell health or even resulting in cell death. Mitigating this apoptotic response can enhance cell survival post-CRISPR editing, which will ultimately increase editing efficiency. In our study, we observed that combining p53 inhibition with pro-survival small molecules yields a homologous recombination rate of over 90% when using CRISPR in human iPSCs. This protocol significantly streamlines the editing process and reduces the time and resources necessary for creating isogenic lines.

Developing a physiologically relevant in vitro model of the respiratory epithelium is critical for understanding lung development and respiratory diseases. Here, we describe a detailed protocol in which the fetal mouse proximal epithelial progenitors were differentiated into 3D airway organoids, which contain terminal-differentiated ciliated cells and basal stem cells. These differentiated airway organoids could constitute an excellent experimental model to elucidate the molecular mechanisms of airway development and epithelial cell fate determination and offer an important tool for establishing pulmonary dysplasia disease in vitro.

Targeted genome editing of human pluripotent stem cells (hPSCs) is critical for basic and translational research and can be achieved with site-specific endonucleases. Cpf1 (CRISPR from Prevotella and Francisella) is a programmable DNA endonuclease with AT-rich PAM sequences. In this protocol, we describe procedures for using a single vector system to deliver Cpf1 and CRISPR RNA (crRNA) for genome editing in hPSCs. This protocol enables indel formation and homologous recombination–mediated precise editing at multiple loci. With the delivery of Cpf1 and a single U6 promoter-driven guide RNA array composed of an AAVS1-targeting and a MAFB-targeting crRNA array, efficient multiplex genome editing at the AAVS1 (knockin) and MAFB (knockout) loci in hPSCs could be achieved in a single experiment. The edited hPSCs expressed pluripotency markers and could differentiate into neurons in vitro. This system also generated INS reporter hPSCs with a 6 kb cassette knockin at the INS locus. The INS reporter cells can differentiate into β-cells that express tdTomato and luciferase, permitting fluorescence-activated cell sorting of hPSC-β-cells. By targeted screening of potential off-target sequences that are most homologous to crRNA sequences, no off-target mutations were detected in any of the tested sequences. This work provides an efficient and flexible system for precise genome editing in mammalian cells including hPSCs with the benefits of less off-target effects.

Primary human intestinal stem cells (ISCs) can be cultured and passaged indefinitely as two-dimensional monolayers grown on soft collagen. Culturing ISCs as monolayers enables easy access to the luminal side for chemical treatments and provides a simpler topology for high-resolution imaging compared to cells cultured as three-dimensional organoids. However, the soft collagen required to support primary ISC growth can pose a challenge for live imaging with an inverted microscope, as the collagen creates a steep meniscus when poured into wells. This may lead to uneven growth toward the center of the well, with cells at the edges often extending beyond the working distance of confocal microscopes. We have engineered a 3D-printed collagen mold that enables the preparation of chamber slides with flat, smooth, and reproducible thin collagen layers. These layers are adequate to support ISC growth while being thin enough to optimize live imaging with an inverted microscope. We present methods for constructing the collagen press, preparing chamber slides with pressed collagen, and plating primary human ISCs for growth and analysis.