Cell Biology

Studying actin-filament assembly into distinct subcellular structures can provide insights into both physiological cellular processes and the mechanisms of disease. However, there are a limited number of tools that can quantify the organization and abundance of different actin structures from confocal microscopy images of cells expressing Lifeact or fixed and stained with phalloidin. Filamentous actin segmentation tool (FAST) is a deep learning model trained with a unique approach of antibody-assisted annotation, resulting in accurate and efficient quantification of distinct classes of actin structures. Here, we detail the protocol for using antibody-assisted annotation to generate datasets that could be applied to train machine learning models. Additionally, we provide step-by-step instructions for applying FAST on phalloidin-stained or live-cell confocal imaging data using our pretrained model. FAST is open source and freely available, with user-friendly notebooks that enable quantification of different classes of actin structure, without the need for structure-specific antibodies. As such, FAST can be a practical tool for researchers investigating the role of cytoskeletal organization in a range of processes.

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.

Aging is associated with progressive accumulation of senescent cells, which contribute to tissue dysfunction and organismal decline. Conventional approaches for assessing cellular senescence, such as histological or immunofluorescence analyses of fixed tissue sections and flow cytometry, require tissue collection, thereby precluding longitudinal in vivo studies. To enable the analysis of cellular senescence in a living vertebrate model, we have previously generated a cdkn1a (p21)-driven GFP reporter line that was established in the transparent klara background of Nothobranchius furzeri. Here, we describe a protocol for in vivo light-sheet microscopy of the reporter activity as readout for senescence-associated cell cycle arrest with single-cell resolution. The procedure involves anesthesia and mounting of fish for stable positioning within the imaging chamber, with particular attention to animal welfare considerations. It further includes the acquisition of three-dimensional image stacks and subsequent image processing. The workflow allows monitoring of GFP-positive cells in intact living killifish at different developmental stages. Although imaging depth remains limited despite organismal transparency, this method provides high-resolution volumetric imaging with minimal phototoxicity and enables analysis of senescence dynamics in a short-lived vertebrate model. It is currently performed as a terminal procedure under approved ethical regulations, but longitudinal imaging would also be possible with additional ethical authorization.

Polyploid hepatocytes are one of the unique features of the liver. Some polyploid hepatocytes have chromosomes in a single nucleus (e.g., 1x4n, 1x8n), while others separate their chromosomes into two nuclei (e.g., 2x2n, 2x4n). In ploidy research, hepatocytes are typically sorted according to their cellular ploidy, revealing their contribution to tumorigenesis and cellular senescence. However, the conventional sorting method fails to distinguish 1x4n from 2x2n, or 1x8n from 2x4n cells, leaving it unclear whether hepatocytes with the same cellular ploidy but different nuclear configurations are identical or phenotypically different. Here, we describe a detailed protocol for fractionating mononucleated and binucleated hepatocytes. First, we present the method for isolating primary mouse hepatocytes and staining them with the DNA dye Hoechst 33342. Flow cytometry is then used to detect fluorescence differences between mononucleated and binucleated hepatocytes. This protocol enables the discrimination of hepatocyte subpopulations with identical cellular ploidy, providing a useful tool to investigate the functional heterogeneity of polyploid hepatocytes.

In the model alga Chlamydomonas reinhardtii, CRISPR (clustered regularly interspaced short palindromic repeat)-based gene editing using Cas (CRISPR-associated) enzymes enables both (a) insertion of large gene cassettes and (b) the creation of knockouts based on the introduction of indels, and specific mutations via mutation-directing oligonucleotides. Owing to the relatively low efficiency of this process, selection markers are frequently used to enrich the candidate pool prior to screening, which typically employs PCR. Unfortunately, few selection markers are available for Chlamydomonas. Furthermore, each marker requires different selection media, and deletion of the selectable marker can be difficult. When multiple successive gene editing steps are required, the use of these markers becomes onerous. The SCREAM (sequential CRISPR via recycling endogenous auxotrophic markers) technique employs an endogenous gene as a marker, the mutation of which can be selected both in the forward (loss of function) and reverse (gain of function) directions. During the first gene editing step, crRNA and mutation-directing oligonucleotides are provided for both the marker and the first target gene (Target 1). Candidates with edited marker genes are selected by loss of marker function, prior to screening for the desired modification of the first target gene. Using a successful candidate, a subsequent gene editing step directs reversion of the mutant marker gene to wild-type status, with candidates being selected on auxotrophic media to detect the regain of function of the auxotrophic marker to wild type (i.e., reversion). Simultaneously, a second target gene modification is produced using Target 2–specific crRNA and oligonucleotides. Revertants, now with a wild-type auxotrophic marker, are then screened for the specific mutation of Target 2. This reversion strategy enables a single selectable marker to be reused indefinitely, facilitating the creation of many successive mutations in a single cell line. As the marker can be completely reconstituted, strains can be created in which only the target gene is altered. Employment of homology-directed repair, using single-stranded oligonucleotides for mutation creation, enables the creation of site-directed mutants, tag insertion, and gene knockouts or reversion, rather than the insertion of large gene cassettes. In this implementation, nitrate reductase is used as the endogenous auxotrophic marker, and the adenine phosphoribosyltransferase gene is used as an example of a target gene.

UFMylation is an evolutionarily conserved ubiquitin-like modification that covalently conjugates UFM1 to lysine residues of substrates via a sequential E1-E2-E3 enzymatic cascade. UFMylation plays a pivotal role in maintaining cellular homeostasis, and its dysregulation is closely linked to multiple major diseases, including malignant tumors, hematopoietic defects, neurodegenerative disorders, and congenital developmental defects, highlighting its important biological significance. However, few substrates of UFMylation have been reported to date, limiting our deep understanding of the mechanistic functions of this modification. This major bottleneck stems from two major technical limitations: the overwhelming abundance of ribosomal protein L26 (RPL26)-UFM1 conjugates masks signals from low-abundance substrates, and conventional methods rely on cumbersome cotransfection of multiple pathway components with poor efficiency and specificity in UFMylated peptides enrichment. To address these challenges, we have developed an effective and specific experimental protocol for UFMylation detection and large-scale substrate identification. This protocol employs CRISPR-Cas9-mediated gene editing to generate UFSP1/UFSP2 double-knockout (UFSP1^KO/UFSP2^KO, DKO) HEK293T cells, which completely abrogate de-UFMylation and thus significantly elevate global protein UFMylation levels upon exogenous introduction of mature UFM1-ΔC2. In addition, exogenous co-expression of the E3 ligase core components UFL1 and DDRGK1 can further improve the sensitivity of substrate detection. This protocol enables large-scale identification of UFMylation substrates with modification sites via high-efficiency enrichment with the K-ε-VG antibody and LC-MS/MS analysis.

Organoids as a drug discovery platform represent an emerging field that continues to refine its tools. NanoBRET (bioluminescence resonance energy transfer) has emerged as a proximity-based and highly sensitive assay to measure protein–protein and protein–ligand interactions. NanoBRET assays were developed and are currently used for 2D cell line experiments. Here, we present the development of the first organoid-compatible Nanoluciferase (Nluc) for 3D model systems. We utilise the Nluc for NanoBRET assays to test drug–target engagement. We describe steps for seeding, transfecting, and replating of mouse colorectal cancer organoids. In addition, we provide detailed procedures for the NanoBRET assay. Various lines of evidence have shown significant difference in drug response between 2D human cell lines and 3D model systems, including patient-derived organoids. Our protocol provides a template for measuring this difference in the context of drug–target engagement.

Satellite cells are adult skeletal muscle stem cells that play essential roles in muscle regeneration. Understanding their behavior is critical for elucidating the mechanisms of muscle repair and advancing muscle regenerative therapies. This requires efficient methods for genetic manipulation in these cells. Retroviral-mediated gene delivery is commonly used for stable transgene expression in immortalized cell lines. However, existing approaches are not optimized for primary satellite cells, often resulting in variable efficiency and inconsistent outcomes. Here, we describe an optimized protocol for satellite cell isolation and culture, as well as retroviral production and infection of primary satellite cells that achieves high transduction efficiency. The satellite cell isolation procedure enriches for myofiber fragments prior to satellite cell release, thereby reducing contamination by non-myogenic cells and improving cell purity. Another key feature of this protocol is the concentration of retroviral particles and their resuspension in satellite cell growth medium prior to infection, which minimizes satellite cell exposure to packaging cell-conditioned medium. Compared to standard approaches, this protocol improves both infection efficiency and reproducibility. It is readily adaptable to a wide range of downstream applications, including microscopies, biochemical assays, and molecular biology analyses.

Organoids are three-dimensional cell structures derived from stem cells that recapitulate the architecture and function of native tissues. Histological analysis of organoids is essential for assessing their structure, cellular composition, and responses to experimental conditions. However, their small size and fragility make standard paraffin embedding workflows difficult. Here, we describe a robust and reproducible protocol for the fixation, paraffin embedding, and sectioning of human organoids, enabling high-quality histological and immunostaining analysis. The method involves direct fixation within the culture matrix and inclusion in HistoGel to prevent organoid loss during processing. The protocol is compatible with hematoxylin–eosin (H&E) staining and multiplex immunofluorescence. Critical steps, troubleshooting, and adaptations for intestinal and cardiac organoids are discussed. This cost-effective and accessible method supports long-term preservation and detailed structural analysis of organoid models.

Seahorse metabolic assays are now widely utilized across numerous fields for performing functional assessments of glycolysis and mitochondrial function in adherent or suspension cell culture samples. Seahorse assays measure extracellular acidification rate (ECAR) and oxygen consumption rate (OCR) as a means of assessing glycolysis and mitochondrial function, respectively. Currently, the vast majority of Seahorse metabolic assays are performed using in vitro samples due to the current established standardized method. However, a uniform approach to assess real-time functional measurements of glycolysis and mitochondrial function in ex vivo tissue samples remains elusive. In particular, this protocol was designed to assess glycolysis in ex vivo murine intestinal samples through ECAR measurements using the Agilent Seahorse XFe24 platform with corresponding Islet Capture microplates and screens. This protocol was developed to provide functional measurements of glycolytic metabolism in murine intestinal tissue samples. This protocol details a method to assess glycolysis in tissue samples and represents the next stage of ex vivo metabolic methods to complement existing standardized in vitro approaches. While this protocol was developed to assess ECAR in ex vivo murine intestinal samples, the same approach can be applied to assessing mitochondrial respiration through measurements of OCR in other tissue types. Overall, this protocol expands the purview of Seahorse metabolic assays through the inclusion of tissue samples and provides the framework to interrogate organ-level metabolism in the context of systemic nutrient metabolism and physiology.