细胞生物学

Human bone marrow organoids provide a tractable three-dimensional platform for modeling hematopoiesis and hematologic disease in a human niche–like context. Here, we describe a stepwise protocol for utilizing human induced pluripotent stem cell (iPSC)-derived bone marrow organoids that support autonomous hematopoiesis for hematopoietic disease modeling, mouse xenograft hematopoiesis, and drug sensitivity testing. The workflow combines embryoid body formation, early mesoderm/angiogenic induction under hypoxia, hemogenic endothelial commitment, maturation within a collagen-containing hydrogel, and subsequent suspension culture as individual organoids. The resulting organoids contain endothelial, stromal, and hematopoietic components and reproduce key structural and cellular features of human marrow. We further describe procedures for engraftment of normal donor- or patient-derived CD34+ cells and implantation of mature organoids under the renal capsule of immunodeficient mice to assess in vivo hematopoietic maintenance. In prior applications of this platform, donor-derived CD34+ cells were shown to engraft within the organoid niche and undergo multilineage differentiation, enabling detection of selective erythroid defects caused by DDX41 deficiency and assessment of therapeutic suppression of JAK2V617F-mutant patient-derived hematopoietic cells in a human marrow–like microenvironment. This protocol, therefore, enables disease modeling, in vivo xenograft assessment, and ex vivo functional analysis of patient-derived hematopoietic cells using relatively small input samples.

Three-dimensional (3D) spheroid models have become essential in cancer biology, drug screening, and tissue engineering. However, their small size, fragile structure, and tendency to disintegrate during routine histoprocessing present persistent technical challenges. Conventional paraffin embedding often results in tissue fragmentation, loss of spatial orientation, and poor section quality, whereas cryosectioning often compromises cellular morphology. Here, we present a robust, cost-effective protocol for preserving and sectioning fragile 3D spheroids, resulting in high-quality histological sections with intact architecture and excellent cellular detail. The method involves optimized handling and embedding procedures that stabilize spheroids during standard formalin fixation, paraffin infiltration, and microtomy, eliminating mechanical distortion and preserving spherical integrity for consistent sectioning. We demonstrate the successful application of this protocol across different tumor spheroids derived from distinct tissue types, including MDA-MB-231 (breast carcinoma), Caco-2 (colorectal adenocarcinoma), and A549 (lung carcinoma) cell lines, with subsequent compatibility with hematoxylin and eosin (H&E) staining protocols. This protocol is compatible with immunohistochemistry (IHC) and immunofluorescence (IF). In our qualitative comparison with conventional methods, we found that our approach effectively limits sample loss, promotes consistent inter-section reproducibility, and preserves fine structural features, such as necrotic cores, proliferative zones, and extracellular matrix components. This protocol provides a reliable, accessible solution for routine histological analysis of fragile 3D spheroids, facilitating more accurate morphological assessment in translational research settings.

The airway epithelium interfaces with the external environment through its apical surface and with the extracellular matrix (ECM) through its basolateral surface. To model this organization in vitro, we developed a decellularized ECM-incorporated apical-out airway organoid (dECM-AoAO) platform in which human bronchial epithelial cells (HBECs) self-assemble around human lung-derived decellularized ECM microparticles (dECM-MPs). This configuration preserves apical-out polarity while enabling direct epithelial–ECM interactions. Here, we describe a protocol for the vacuum filtration and quantification of dECM-MPs, the generation of dECM-AoAOs, and ultimately, whole-mount immunofluorescence staining for organoid characterization.

Skin models play critical roles in understanding disease mechanisms and advancing therapeutic development. However, conventional systems based on 2D cell cultures, in vivo animal models, and ex vivo tissue explants are limited by insufficient physiological complexity, interspecies differences, and restricted accessibility, respectively. Advances in biofabrication technologies have enabled the engineering of 3D skin equivalents that better balance biological complexity and experimental scalability. Here, we present a biofabrication protocol inspired by the regenerative processes of wound healing to construct vascularized 3D organotypic skin models in a stepwise manner. The approach integrates bioprinting for precise spatial organization of cellular compartments with guided cell self-organization to achieve native-like tissue complexity and heterogeneity. Through a programmable culture strategy, tissue maturation proceeds sequentially through keratinocyte proliferation and collective migration, microchannel endothelialization, basal-to-suprabasal differentiation, and progressive extracellular matrix remodeling within a fibrin-based scaffold. The resulting tissue constructs comprise stratified epidermal layers positioned atop a vascularized, fibroblast-remodeled dermal matrix. Beyond reproducing key structural features of human skin, this protocol recapitulates cellular processes associated with tissue regeneration, providing a dynamic platform for investigating disease pathogenesis, progression, and therapeutic responses.

Tumor mechanical microenvironment, particularly extracellular matrix stiffness, plays a critical role in regulating cancer cell behavior, including proliferation, quiescence, and drug resistance. Conventional 2D culture or stiff 3D scaffolds fail to recapitulate the physiological soft (normal) or pathologically stiff (tumoral) mechanical niches. Here, we present a detailed protocol for establishing a tunable 3D tumor spheroid culture system using sodium alginate–based hydrogels crosslinked with calcium ions at different concentrations to achieve soft or stiff conditions that mimic normal colon and colorectal cancer tissues, respectively. We describe the step-by-step procedures for fabricating stiffness-tunable hydrogels, culturing colorectal cancer spheroids, releasing spheroids for downstream analysis, and performing immunohistochemical staining on intact spheroids. This protocol enables the reproducible investigation of mechanosensitive pathways and drug resistance mechanisms in a physiologically relevant 3D context.

In vitro cytotoxicity assessments frequently rely on staining-based methods that indirectly estimate viable cell numbers. A major limitation of many such techniques is their endpoint nature, requiring cell lysis or irreversible processing that precludes longitudinal monitoring of cellular responses following treatment. An ideal assay for evaluating cell viability and proliferation should be simple, rapid, cost-effective, reproducible, and highly sensitive, while also enabling accurate quantification with minimal interference from test compounds. The resazurin reduction assay satisfies these criteria, offering a sensitive and economical alternative to conventional tetrazolium-based methods. Although both assay types depend on the metabolic reduction of a dye by viable cells, they differ mechanistically. Tetrazolium salts (e.g., MTT) are reduced by cellular dehydrogenases to insoluble formazan crystals that require solubilization before detection. In contrast, resazurin-a cell-permeable, non-fluorescent blue dye-is reduced to resorufin, a highly fluorescent compound detectable without additional processing steps. This property renders the resazurin assay broadly applicable to viability testing in eukaryotic cells cultured in both 2D and 3D formats, as well as in bacterial systems. Here, we present a resazurin-based reduction assay across diverse experimental models, emphasizing its practicality, reproducibility, and adaptability for real-time viability monitoring.

Breast cancer (BC) is the most frequently diagnosed malignancy in women and a leading cause of cancer-related mortality worldwide. Current clinical management relies on molecular classification—based on estrogen receptor (ER), progesterone receptor (PR), HER2, and Ki67 expression—to guide prognosis and therapy. Triple-negative breast cancer (TNBC), which lacks ER, PR, and HER2 expression, represents 15%–20% of cases and is characterized by aggressive behavior, early recurrence, and a paucity of targeted treatment options. These challenges underscore the urgent need for improved preclinical models that better recapitulate tumor biology to accelerate therapeutic discovery. While conventional monolayer (2D) cultures have contributed significantly to cancer research, they fail to mimic critical features of the three-dimensional (3D) tumor microenvironment (TME), thereby limiting clinical translation. To address this gap, 3D spheroid models have emerged as a powerful intermediary, more accurately replicating in vivo conditions such as cell–cell and cell–matrix interactions, nutrient and oxygen gradients, and the development of hypoxic cores. These features make spheroids a physiologically relevant platform for studying complex processes like metastasis, drug resistance, and treatment response. Here, we present a robust, simple, and cost-effective protocol for generating uniform 3D spheroids. Our method enables consistent monitoring of spheroid formation and growth over time, with quantitative, image-based size analysis to ensure reproducibility and scalability. Designed for flexibility, the protocol is broadly applicable across diverse cell types, effectively bridging the gap between traditional 2D cultures and complex in vivo studies. By providing an accessible and reliable model of the 3D TME, this protocol opens new avenues for high-throughput drug screening, mechanistic studies of tumor progression, and the advancement of personalized medicine strategies in breast cancer and beyond.

Nowadays, the use of 3D cultures (organoids) is considered a valuable experimental tool to model physiological and pathological conditions of organs and tissues. Organoids, retaining cellular heterogeneity with the presence of stem, progenitor, and differentiated cells, allow the faithful in vitro reproduction of structures resembling the original tissue. In this context, the growth of endometrial organoids allows the generation of 3D cultures characterized by a hollow lumen, secretory activity, and apicobasal polarity and displaying phenotypical modification in response to hormone stimulation. However, a limitation in currently used models is the absence of stromal cells in their structure; as a result, they miss epithelial–stromal interactions, which are crucial in endometrial physiology. We developed a novel 3D model to generate endometrial organoids grown in floating Matrigel^TM droplets in the presence of standard culture medium. From a structural point of view, these novel floating 3D cultures develop as gland-like structures constituted by epithelial cells organized around a central lumen and retain the expression of endometrial and decidual genes, like previously published organoids, although with a phenotype resembling hormonally differentiated structures. Importantly, floating organoids retain stromal cells which grow in close contact with the epithelial cells, localized within the internal or external portion of the organoid structure. In summary, we present a simple and rapid model for generating 3D endometrial organoids that preserve epithelial–stromal cell interactions, promoting the formation of differentiated organoids and enabling the study of reciprocal modulation between epithelium and stroma.

Musculoskeletal pathologies present challenges in athletic horses, often leading to functional impairment. The slow or limited regenerative capacity of bone, joint, and tendon/ligament injuries, coupled with the limitations of conventional treatments, highlights the need for innovative therapies such as ortho-biologics and mesenchymal stem/stroma cells. Traditional 2D cell culture systems with fetal bovine serum (FBS) fail to replicate the complexity of the in vivo environment, whereas 3D cultures more accurately mimic native tissue architecture and cell–cell interactions. This study describes a novel method for isolating muscle-derived progenitor cells in a 3D environment using an autologous plasma-based gel and an innovative cell retrieval solution. The cultured cells exhibit immunomodulatory effects on T lymphocytes, trilineage differentiation potential, and immunophenotypic characteristics consistent with conventional mesenchymal stem/stromal cells. This streamlined 3D culture technique offers a promising platform for generating minimally manipulated autologous cell products tailored for equine regenerative medicine.

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.