Understanding radiation-induced tissue modifications and cell death within the complex cellular architecture of the brain requires experimental systems that preserve tissue integrity and intercellular crosstalk. At the same time, microglial cells as primary regulators of primary and secondary inflammation in the brain are difficult to study in single cell cultures and it is crucial to investigate their behaviour in their complex tissue microenvironment. Rodent organotypic hippocampal slice cultures (OHSCs) and human patient-derived brain tumor cultures (PDTCs) offer physiologically relevant platforms for investigating the differential effects of X-ray and heavy-ion irradiation on glial as well as immune cell populations. Both models retain native cytoarchitecture, extracellular matrix composition, and functional cell-cell interactions, making them uniquely suited to study the spatial and temporal dynamics of inflammation or cell death in a multicellular context. Here, we present a standardized methodology for preparing, irradiating, and assessing PDTCs, employing multimodal readouts as cytokine measurement of the supernatant, cell death, microglia morphology and function. Novel molecular analyses are possible upon certain considerations and may alter our understanding of species differences and help to distinguish cell death pathways to attribute damage to specific cell types. Standardization of these protocols across model systems is essential for generating reproducible, comparative data while preserving the inherent complexity that underlies intercellular communication and coordinated responses to injury. This approach enables translationally relevant insight into the mechanisms of radiation-induced brain injury to develop strategies for neuroprotection and treatment in clinical settings.
The detection and measurement of lipid reactive oxygen species (ROS) is a critical method widely applied in the study of ferroptosis, a recently defined form of iron-dependent programmed cell death characterized by excessive lipid peroxidation and subsequent membrane damage. As ferroptosis research expands into various pathophysiological processes and human diseases, including cancer, immunity and inflammation, and tissue injuries, there is an increasing need for robust methods to measure lipid ROS in heterogenous cell samples, such as in deep immune cell profiling. BODIPY™ C11 581/591 is a gold-standard ratiometric fluorescent dye that reports the levels of lipid ROS, and has been used in both mechanistic and applied studies demonstrating ferroptosis regulation across multiple cell types. However, its use has largely been confined to traditional flow cytometry due to its extensive spillover into multiple emission channels and its ratiometric properties, which complicate its application in spectral flow cytometry. Herein, we present solutions to these challenges and a comprehensive workflow for the optimization, unmixing, staining, and analysis required to multiplex lipid ROS detection using BODIPY™ C11 581/591 with high-parameter spectral flow cytometry panels. Our unmixing strategy represents a novel method for separating the fluorescent spectra of two forms of a ratiometric dye, which may also be applicable to other ratiometric dyes in spectral flow cytometry.
Macrophages are key innate immune effector cells capable of responding to microbial factors such as lipopolysaccharide (LPS) through activation of inflammatory pathways. However, repeated LPS exposure can lead to endotoxin tolerance, a reprogrammed state where macrophages suppress pro-inflammatory cytokine production while preserving antimicrobial functions. The reduction in inflammatory responses during endotoxin tolerance serves to preserve tissue integrity and macrophage viability, although it can also result in immunosuppression, as observed in sepsis. An enhanced understanding of the molecular mechanisms which govern endotoxin tolerance in macrophages could unlock new ways to modulate innate immunity. Here we provide detailed protocols for inducing and measuring tolerance in murine bone marrow-derived macrophages (BMDMs) and human PMA-differentiated THP-1 macrophages, using ELISA-based quantification of IL-1β and IL-6 as functional readouts. These methods provide robust in vitro systems to study endotoxin tolerance.
Apoptotic bodies (ApoBDs) are a type of extracellular vesicle (EV) generated during the final stages of apoptosis. These vesicles were traditionally regarded as apoptotic cellular waste. More recently, they have begun to be recognized as mediators of intercellular communication via the delivery of diverse factors, with emerging roles in various physiological processes including immune regulation, cancer progression, and tissue repair. However, the lack of standardized methods for isolating and characterizing ApoBDs has limited both their study and potential application. Here, we present simple, efficient, and reproducible protocols for ApoBD isolation, characterization, and application to downstream experimental approaches. This protocol includes a two-step differential centrifugation to isolate ApoBDs and flow cytometry-based characterization, as well as optional steps such as caspase inhibitor-based modulation of ApoBD release and fluorescent labeling of ApoBDs for their tracking. Furthermore, we provide methods to evaluate ApoBD uptake by recipient cells, enabling subsequent downstream functional studies. These approaches promote the reproducibility and standardization of ApoBD research, providing a foundation to deepen our understanding of their functional roles and advance future clinical applications.
Regulated cell death mediated by dedicated molecular machines, known as programmed cell death, plays important roles in health and disease. Understanding the mechanisms of cell death is crucial for elucidating the control of cellular homeostasis and developing therapies for related diseases. Despite extensive research efforts spanning decades, many aspects of cell death mechanisms remain elusive, highlighting the need for continued exploration. Here, we describe how to identify novel regulators involved in cell death pathways using a genome-wide screening approach.
Cellular senescence is a stable form of cell cycle arrest triggered by diverse stressors such as telomere shortening, oncogene activation, DNA damage, and chemotherapy. While senescence acts as a tumor-suppressive mechanism, the long-term accumulation of senescent cells contributes to chronic inflammation, tissue dysfunction, and age-related diseases, largely through the senescence-associated secretory phenotype (SASP). Senescent cells are characterized by increased expression of p16INK4a and p21CIP1, enlarged morphology, and resistance to apoptosis. This apoptotic resistance is mainly driven by the upregulation of anti-apoptotic BCL-2 family of proteins, including BCL-2, BCL-xL, and MCL-1. Senolytics - drugs that selectively induce apoptosis in senescent cells - target these survival pathways to promote senescent cell clearance. BH3 profiling is a functional assay that assesses mitochondrial apoptotic priming by exposing permeabilized cells to BH3-domain peptides and measuring cytochrome c release. This technique allows the identification of anti-apoptotic dependencies in senescent cells and can be used in combination with senescence markers (p16 and p21) via flow cytometry to pinpoint populations with enhanced apoptotic resistance. Understanding the apoptotic landscape of senescent cells is critical for optimizing senolytic strategies and improving therapeutic outcomes, particularly in contexts such as cancer and therapy-induced senescence.
Inflammation and cell death are central to both chronic inflammatory disorders and cancer progression, influencing tissue homeostasis, therapeutic response, and resistance. Effective preclinical models that preserve the human microenvironment are required to study these processes, guide personalized therapy or to test pharmacologically active substances. We developed patient-derived tissue slice cultures from endoscopic specimens (ePDTC) such as gastrointestinal and pulmonary cancers, as well as inflamed mucosa from patients with Inflammatory bowel disease (IBD). These tissue cultures preserve epithelial, stromal, and immune compartments, enabling analysis of inflammatory signalling and cell death pathways while allowing parallel pharmacological testing. PDTCs maintained tissue cytoarchitecture and cell-cell interactions, permitting within-patient drug response profiling and capturing inter- and intra-patient heterogeneity. This enabled mechanistic investigation of inflammation-driven resistance and individualized assessment of chemotherapies, immunotherapies, and anti-inflammatory agents. Here, we describe a protocol for establishing ePDTC from diverse anatomical sites to provide a translational platform to study immune regulation and therapy response in human cancers and inflammatory diseases, supporting development of personalized treatment strategies.
The PIDDosome multiprotein complex is formed by PIDD1 and RAIDD (alias CRADD) and serves as activation platform for caspase-2. One of the best characterized triggers for PIDDosome activation is the presence of extra centrosomes, which are frequently observed in cancer and in naturally polyploid tissues such as liver and heart. Depending on the cell type, activated caspase-2 can (1) cleave MDM2, thereby triggering a p53 response, or (2) cleave the BH3-only protein BID to induce apoptosis. Here, we describe our biochemical and cell biological tools to study the PIDDosome and caspase-2 activity. These include methods to experimentally induce PIDDosome formation in cells, as well as techniques to monitor pathway activity via protein and mRNA expression analysis, and flow cytometry. Moreover, we describe how to follow centrosome maturation for PIDDosome activation using immunofluorescence microscopy.
Regulated cell death (RCD) is a fundamental biological process that ensures tissue homeostasis, mediates stress responses, and contributes to diverse pathological conditions. The budding yeast Saccharomyces cerevisiae has emerged as a valuable model organism for the study of RCD, providing evolutionary insights into conserved molecular pathways and enabling the systematic dissection of cell death mechanisms in a genetically tractable system. Upon a lethal stimulus, yeast populations may segregate into three major subtypes: primary necrotic cells with disrupted plasma membranes, early apoptotic cells exhibiting morphological markers of apoptosis, and late apoptotic (or secondary necrotic) cells that display both apoptotic and necrotic features. Discriminating between these subpopulations is essential for accurate interpretation of RCD dynamics and has been facilitated by the transition from fluorescence microscopy to flow cytometry, which allows rapid, quantitative, and high-throughput analysis. This manuscript provides a methodological framework for the flow cytometric identification and quantification of these distinct subpopulations of dead or dying yeast, enabling reproducible and detailed assessment of cell death heterogeneity.
The Lysosomal Galectin Puncta Assay is a microscopy-based technique able to detect even minor lysosomal leakage with high sensitivity. This protocol describes the detection of galectin puncta as markers of lysosomal membrane permeabilization, a process that relies on the high-affinity binding of the cytosolic galectins to the luminal glycans exposed on damaged lysosomes. Compared to traditional methods, the Galectin Puncta Assay offers high sensitivity, detects subtle lysosomal leakage, and enables analysis at single-lysosome level. Here, we provide a step-by-step protocol for this assay, covering sample preparation, immunostaining, imaging and image quantification.
Pannexin 1 (PANX1) is a ubiquitously expressed plasma membrane channel that plays an important role in maintaining cellular homeostasis through the release of small signalling metabolites. Under cell death settings, PANX1 channels can become activated by caspase-3/7 cleavage. This leads to the release of chemotactic 'find-me' signals and anti-inflammatory 'good-bye' signals. Additionally, caspase-activated PANX1 channels can regulate the fragmentation of dying cells. Due to the various cellular pathways that PANX1 channels are involved in, this membrane channel has emerged as an attractive target for novel therapeutics. To assess the effectiveness of pharmacological inhibition on caspase-activated PANX1 channels, the cell-impermeable nucleic acid-binding dye, TO-PRO-3 can be used due to its ability to be selectively taken up into apoptotic cells through caspase-activated PANX1 channels. Here we describe a robust flow cytometry-based protocol utilising Annexin A5 and TO-PRO-3 staining to accurately monitor apoptotic cell death and PANX1 channel activity. This approach can be easily adapted to investigate pharmacological compounds that have the potential to inhibit caspase-activated PANX1 channels.
