Cell Biology

Adipose tissue macrophages (ATMs) critically influence obesity-induced inflammation and metabolic dysfunction. Recent studies identified distinct ATM subsets characterized by markers such as CD11c, CD9, and Trem2, associated with pro-inflammatory and metabolically activated states. This protocol outlines a detailed, reproducible methodology for isolating, characterizing, and sorting these ATM subsets from murine epididymal white adipose tissue (eWAT) using multicolor flow cytometry. Key steps include stromal vascular fraction (SVF) isolation, immunophenotyping, sequential gating strategies, and fluorescence-activated cell sorting (FACS) for downstream gene expression analysis. The protocol was validated in diet-induced obese (DIO) mice treated with the IRE1 RNase inhibitor STF-083010, demonstrating its utility for studying ATMs in the context of obesity and metabolic disease.

Inherited germline variants are now recognized as important contributors to hematologic myeloid malignancies, but their reliable detection depends on obtaining uncontaminated germline DNA. In solid tumors, peripheral blood remains free of tumor cells and therefore serves as a standard source for germline testing. In contrast, peripheral blood often contains neoplastic or clonally mutated cells in hematologic malignancies, making it impossible to distinguish somatic from germline variants. This unique challenge necessitates using an alternative, non-hematopoietic tissue source for accurate germline assessment in patients with hematologic myeloid malignancies. Cultured skin fibroblasts derived from punch biopsies have long been considered the gold standard for this purpose. Nevertheless, most existing protocols are optimized for research settings and lack detailed, patient-centric workflows for routine clinical use. Addressing this translational gap, we present a robust, enzyme-free protocol for culturing dermal fibroblasts from skin punch biopsies collected at the bedside during routine bone marrow procedures. The method details practical bedside collection, sterile transport, mechanical dissection without enzymatic digestion, plating strategy, culture expansion, and high-yield DNA isolation with validated purity. By integrating this standardized approach into routine hematopathology workflows, the protocol ensures reliable germline material with minimal patient discomfort and a turnaround time suitable for clinical diagnostics.

Intestinal organoids are generated from intestinal epithelial stem cells, forming 3D mini-guts that are often used as an in vitro model to evaluate and manipulate the regenerative capacities of intestinal epithelial stem cells. Plating 3D organoids on different substrates transforms organoids into 2D monolayers, which self-organize to form crypt-like regions (which contain stem cells and transit amplifying cells) and villus-like regions (which contain differentiated cells). This “open lumen” organization facilitates multiple biochemical and biomechanical studies that are otherwise complex in 3D organoids, such as drug applications to the cell’s apical side or precise control over substrate protein composition or substrate stiffness. Here, we describe a protocol to generate homogenous intestinal monolayers from single-cell intestinal organoid suspension, resulting in de novo crypt formation. Our protocol results in higher viability of intestinal cells, allowing successful monolayer formation.

Crypts at the base of intestinal villi contain intestinal stem cells (ISCs) and Paneth cells, the latter of which work as niche cells for ISCs. When isolated and cultured in the presence of specific growth factors, crypts give rise to self-renewing 3D structures called organoids that are highly similar to the crypt-villus structure of the small intestine. However, the organoid culture from whole crypts does not allow investigators to determine the contribution of their individual components, namely ISCs and Paneth cells, to organoid formation efficiency. Here, we describe the method to isolate Paneth cells and ISCs by flow cytometry and co-culture them to form organoids. This approach allows the determination of the contribution of Paneth cells or ISCs to organoid formation and provides a novel tool to analyze the function of Paneth cells, the main component of the intestinal stem cell niche.

The female reproductive tract is comprised of different regions, each with distinctive physiological characteristics. One of them is the fallopian tubes, which are vital for human reproductive health and success. The ability to model their function and physiology is of utmost importance. So far, in vitro models have been based on a few immortalized or cancer cell lines derived from fallopian tube cells that lacked differentiated, specialized cell types and did not allow for the study of cancer initiation due to their implicit biases. Organoids, in contrast, overcome these limitations and provide an advanced, three-dimensional system for the study of healthy fallopian tube physiology and pathology. Fallopian tube organoids are comprised of epithelial progenitors that can be enriched using chemical or hormonal treatment into the different cell types that are found in the in vivo tissue, namely detyrosinated-tubulin-positive ciliated cells or paired-box protein 8 (PAX8)-positive secretory cells. This protocol provides a step-by-step guide for the establishment and maintenance of a long-term culture of organoids from healthy human fallopian tube tissue. The organoid model described here closely mimics the in vivo physiology and anatomy of human fallopian tube epithelium and provides a comprehensive basis for future studies on its underlying molecular characteristics and possible pathology.

Regulatory T cells (Tregs) are essential for maintaining immune balance by controlling the activation and expansion of other immune cells. Conventional suppression assays often rely on co-culturing purified cell populations, which limits multiplexed phenotyping and physiological relevance. This protocol describes a high-dimensional, single-cell assay for profiling Treg-mediated suppression within a peripheral blood mononuclear cell (PBMC) system. Tregs are first isolated by cell sorting and then reintroduced into autologous PBMCs at defined ratios. A 52-marker mass cytometry (CyTOF) panel is used to quantify cell division and phenotypic responses across multiple immune subsets. This approach allows for integrated analysis of Treg function with broad compatibility for patient profiling and drug evaluation.

The skin microbiome, a diverse community of microorganisms, plays a crucial role in maintaining skin health and homeostasis. Traditional studies have relied on two-dimensional (2D) models, which fail to recreate the complex three-dimensional (3D) architecture and cellular interactions of in vivo human skin, and animal models, which have species-specific physiology and accompanying ethical concerns. Consequently, both types of models fall short in accurately replicating skin physiology and understanding its complex microbial interactions. Three-dimensional bioprinting, an advanced tissue engineering technology, addresses these limitations by creating custom-designed tissue scaffolds using biomaterial-based bioinks containing living cells. This approach provides a more physiologically relevant 3D structure and microenvironment, allowing the incorporation of microbial communities to better reflect in vivo conditions. Here, we present a protocol for 3D bioprinting an in vitro skin infection model by co-culturing human keratinocytes and dermal fibroblasts in a high-viscosity, fibrin-based bioink to mimic the dermis and epidermis. The bioprinted skin tissue was co-infected with Staphylococcus aureus and Staphylococcus epidermidis to mimic bacterial skin disease. Bacterial survival was assessed through colony-forming unit enumeration. By incorporating bacteria, this protocol offers the potential to serve as a more representative in vivo 3D bioprinted skin infection model, providing a platform to study host–microbe interactions, immune responses, and the development of antimicrobial therapeutics.

Well-differentiated airway epithelial cultures are commonly used to study airway stem cell lineages, ion and fluid transport, respiratory virus infection and replication, and disease mechanisms in vitro. This culture model involves the isolation and expansion of airway stem cells followed by their differentiation at an air–liquid interface (ALI), a process that has been previously documented in humans and mice. Domestic ferrets (Mustela putorius furo) have gained considerable importance in respiratory disease research due to their notable susceptibility to these conditions and their anatomical similarities to humans. Here, we present a comprehensive description of the isolation and culture of stem/progenitor cells from the ferret airway, along with a protocol for their differentiation at the ALI. Our findings have demonstrated that this ferret culture system not only supports the differentiation of the predominant airway epithelial cell types but also facilitates the generation of rare airway epithelial subpopulations, including pulmonary ionocytes, tuft cells, and pulmonary neuroendocrine cells. Additionally, we provide a detailed procedure for measuring transepithelial ion transport relevant to airway diseases, particularly cystic fibrosis. The ability to isolate and culture ferret airway stem cells, combined with ALI differentiation and functional assessment of transepithelial ion transport, offers a powerful platform for evaluating genetic and pharmacologic interventions related to cystic fibrosis.

The fatal motor neuron (MN) disease amyotrophic lateral sclerosis (ALS) is characterized by progressive degeneration of the phrenic MNs (phMNs) controlling the activity of the diaphragm, leading to death by respiratory failure. Human experimental models to study phMNs are lacking, hindering the understanding of the mechanisms of phMN degeneration in ALS. Here, we describe a protocol to derive phrenic-like MNs from human induced pluripotent stem cells (hiPSC-phMNs) within 30 days. During spinal cord development, phMNs emerge from specific MN progenitors located in the dorsalmost MN progenitor (pMN) domain at cervical levels, under the control of a ventral-to-dorsal gradient of Sonic hedgehog (SHH) signaling and a rostro-caudal gradient of retinoic acid (RA). The method presented here uses optimized concentrations of RA and the SHH agonist purmorphamine, followed by fluorescence-activated cell sorting (FACS) of the resulting MN progenitor cells (MNPCs) based on a cell-surface protein (IGDCC3) enriched in hiPSC-phMNs. The resulting cultures are highly enriched in MNs expressing typical phMN markers. This protocol enables the generation of hiPSC-phMNs and is highly reproducible using several hiPSC lines, offering a disease-relevant system to study mechanisms of respiratory MN dysfunction. While the protocol has been validated in the context of ALS research, it can be adopted to study human phrenic MNs in other research fields where these neurons are of interest.

Glomerular diseases characterized by injury to post-mitotic epithelial cells called podocytes are a leading cause of chronic kidney disease. Yet, isolating podocytes from the kidney for transcriptomic, proteomic, and metabolomic studies has been a major technical challenge. Protocols utilizing glomerular sieving and laser capture methods are of limited use because they are not podocyte-specific but instead capture all four glomerular cell types. Here, we present a magnetic-activated cell sorting (MACS) method where podocytes are isolated from digested whole kidneys using antibodies specific to extracellular antigens on podocytes. Using microbeaded secondary antibodies binding to the podocyte-specific primary antibodies allows sorting of the podocytes using a magnet. This podocyte-only cell fraction is a unique source of in vivo–derived cells for molecular and cellular experiments.