STAR Protocols

This protocol describes a Langendorff-based method for isolating intact adult mouse ventricular myocytes using syringe pump-driven perfusion. The approach retains the key physiological advantage of the conventional Langendorff technique, continuous retrograde coronary perfusion, while simplifying the overall setup. By combining retrograde aortic perfusion with widely available laboratory equipment, the method provides an accessible alternative to traditional Langendorff systems. A precision syringe pump connected to an in-line heater is used to deliver temperature-controlled, constant-flow perfusion during enzymatic digestion. In contrast to gravity-driven constant-pressure systems, constant-flow perfusion maintains stable enzyme delivery despite changes in coronary resistance that occur during tissue digestion. Use of an inline heater allows precise, rapid temperature-controlled delivery, avoiding the complexity, leak risk, thermal lag, and contamination susceptibility associated with traditional water-jacketed systems. Our setup reduces variability in perfusion rate and minimizes susceptibility to occlusion, flow interruption, or compliance-related artifacts, enhancing reproducibility. The method consistently yields adult ventricular myocytes with high viability (>70% rod-shaped, calcium-tolerant), enabling a broad range of functional analyses including electrophysiology, contractile performance and calcium handling. Step-by-step instructions, troubleshooting guidance, and anticipated outcomes are provided to facilitate adoption in laboratories without dedicated isolated-heart perfusion infrastructure. Key FeaturesO_LISimplified Langendorff-based mouse cardiomyocyte isolation method that eliminates the need for specialized perfusion rigs. C_LIO_LISyringe pump-driven constant-flow perfusion combined with inline temperature control improves reproducibility by ensuring stable enzyme delivery and precise temperature regulation. C_LIO_LIGenerates high-yield, calcium-tolerant adult mouse ventricular myocytes suitable for functional studies. C_LI Graphical Overview O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=190 SRC="FIGDIR/small/718810v1_ufig1.gif" ALT="Figure 1"> View larger version (63K): org.highwire.dtl.DTLVardef@15061cdorg.highwire.dtl.DTLVardef@44fd48org.highwire.dtl.DTLVardef@1509285org.highwire.dtl.DTLVardef@c362a4_HPS_FORMAT_FIGEXP M_FIG Graphical overview of the simplified Langendorff-based mouse cardiac myocyte isolation protocol. C_FIG

The translation of mRNA into protein is tightly regulated by both cellular trans-factors and cis-regulatory elements encoded within transcripts. Although transcript fate can be measured by transcript abundance or translation efficiency, separating the contribution of each individual cis-element within a single transcript is an ongoing challenge. Current massively parallel reporter assay (MPRAs) approaches enable systematic interrogation of cis-regulatory elements that control transcript stability, but translation-focused MPRAs remain technically limited and often inaccessible. Here we present Nascent Peptide Translating Ribosome Affinity Purification (NaP-TRAP), a reporter-based approach that simultaneously measures translation and mRNA abundance. Unlike previous methods, NaP-TRAP captures translation directly through the immunoprecipitation of epitope-tagged nascent peptide chains, providing instantaneous, frame-specific readouts without specialized instrumentation. The method is highly scalable from single reporters to complex libraries, and adaptable across in vivo and in vitro systems. NaP-TRAP is versatile, allowing assessment of cis-regulatory impact of elements distributed throughout the mRNA, from cap-to-tail. This protocol covers experimental design, reporter construction, sample processing, and computational analysis for both low- and high-throughput applications. Bench work can be completed in 4- 5 days, with qPCR-based readouts requiring only basic Excel skills for data processing. Sequencing-based readouts require skills in command-line tools and Python scripting and add an additional 2-3 days. NaP-TRAP thus offers an accessible, robust, and quantitative platform to decode the regulatory logic of mRNA translation and stability in diverse biological contexts. Basic Protocol 1Design, assembly, and synthesis of NaP-TRAP reporter libraries. Support Protocol 1Design, assembly, and synthesis of NaP-TRAP individual reporters and spike-ins. Basic Protocol 2NaP-TRAP delivery by micro-injection in zebrafish embryos. Alternate Protocol 1NaP-TRAP delivery by transfection in cultured mammalian cells. Basic Protocol 3NaP-TRAP pulldown and RNA extraction. Basic Protocol 4Preparation of NaP-TRAP cDNA Sequencing Libraries. Alternate Protocol 2NaP-TRAP-qPCR module for low-cost validation. Basic Protocol 5Computational analysis of NaP-TRAP MPRA data.

Efficient protein synthesis in eukaryotic cells typically requires a 5' cap structure on messenger RNAs (mRNAs). However, under stress conditions or in viral infection, translation can also occur independently of the cap via internal ribosomal entry sites (IRES). IRES elements are therefore key regulators of protein expression in both viral and cellular contexts. Here we describe a cell-free protocol to quantitatively assess IRES-mediated translation using wheat germ extract (WGE) and a firefly luciferase (FLuc) reporter. The protocol includes template preparation, RNA synthesis and luminescence measurement following in vitro translation in WGE. This method enables rapid and robust comparison of IRES activity under controlled conditions and can additionally be applied to evaluate mRNA modifications designed to enhance translation efficiency. Key featuresO_LIStringent in vitro workflow from DNA template preparation through RNA synthesis and protein synthesis to reporter readout, including quality controls. C_LIO_LIEvaluation of IRES-driven translation suitable for testing combinations of IRES and CDS. C_LIO_LItranslation analysis without radioactive labeling. C_LI Graphical overview O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=89 SRC="FIGDIR/small/716985v1_ufig1.gif" ALT="Figure 1"> View larger version (24K): org.highwire.dtl.DTLVardef@1457b00org.highwire.dtl.DTLVardef@8e7405org.highwire.dtl.DTLVardef@6303eforg.highwire.dtl.DTLVardef@974d71_HPS_FORMAT_FIGEXP M_FIG C_FIG Graphical AbstractPipeline for the production and evaluation of IRES-firefly luciferase constructs using wheat germ extract. (1-4) Preparation: IRES-firefly luciferase constructs are amplified in E. coli and isolated from bacterial cells. Plasmids are linearized to prepare for in vitro transcription. (5-6) Transcript synthesis and verification: In vitro transcription is followed by electrophoretic validation to confirm integrity and correct molecular weight. (7-8) Translation and detection: Translation is executed in wheat germ extract and quantified by measuring reporter activity in a luminometer.

This protocol enables rapid CRISPR-Cas9 genome editing in Saccharomyces cerevisiae by replacing restriction/ligation guide cloning with PCR-based protospacer installation and seamless plasmid recircularization. It describes in silico HDR donor and SgRNA design, install guide sequences into cas9 plasmid by PCR and seamless assembly, plasmid cloning and sequence verification in E. coli, and LiAc/PEG co-transformation of yeast with Cas9-sgRNA plasmid plus HDR donor. The workflow selects yeast colonies on G418 and confirms edits by PCR and sequencing.

Nuclei isolation from myelin-rich adult mouse brain regions remains challenging for single-nucleus RNA sequencing because myelin and debris can reduce nuclei quality. We describe an optimized protocol for mouse hippocampi and cerebella using tube-and-pestle homogenization and low-volume sucrose-gradient pelleting with a standard benchtop centrifuge, with optional magnetic enrichment of nuclei to reduce debris/non-nuclear carryover. Under the tested conditions, the workflow produces intact, debris-reduced nuclei and supports downstream 10x Genomics Flex and PARSE WT library preparation. Graphical abstract O_FIG O_LINKSMALLFIG WIDTH=196 HEIGHT=200 SRC="FIGDIR/small/716374v1_ufig1.gif" ALT="Figure 1"> View larger version (35K): org.highwire.dtl.DTLVardef@ccbd87org.highwire.dtl.DTLVardef@1aef4bcorg.highwire.dtl.DTLVardef@14569a8org.highwire.dtl.DTLVardef@1bc261_HPS_FORMAT_FIGEXP M_FIG C_FIG HighlightsO_LIBenchtop sucrose-gradient pelleting enables rapid nuclei purification from myelin-rich adult mouse brain C_LIO_LIScales across tissue inputs (e.g., hippocampus [~]15-20 mg; cerebellum [~]50-70 mg) without ultracentrifugation or 15 mL gradients C_LIO_LIMagnetic enrichment as the recommended final cleanup step further reduces myelin/debris carryover and is compatible with 10x Flex and PARSE WT workflows. C_LI

RNase R is a processive 3 to 5 exoribonuclease that degrades a broad array of linear RNA species while preserving RNA lariats and circular RNAs (circRNA). In recent years, this enzyme has become pivotal for the field of circRNA research, serving as a key step for circRNA enrichment, purification, and identification. Despite this growing importance, the effects of mutations and truncations in RNase R have been incompletely studied. We make several point mutations and assay their effects on the ability of RNase R to bind and/or degrade RNA substrates. Our data show that selected active site mutations have varying effects on RNA binding and degradation. Furthermore, the increasing interest in circRNA-based RNA therapeutic platforms highlights an urgent need for RNase R in RNA molecular biology labs. However, the substantial cost of commercial RNase R remains a bottleneck, particularly for large-scale studies or the development of circRNA-based technologies. In this protocol, we offer a solution to that problem, namely a more accessible and cost-effective means of purifying high-quality and low-cost RNase R. We provide a highly detailed yet simplified, high-yield protocol that produces recombinant RNase R from Escherichia coli. The method uses a single-step Ni-NTA affinity chromatography procedure without proteolytic tag removal and is optimized for entry-level FPLC systems such as the AKTA Start, ensuring that high-purity enzyme production does not require specialized, high-end instrumentation. A second key feature is the establishment of an optimized reaction framework, including specific buffer compositions and defined enzyme-to-substrate ratios for the purified RNase R. The protocol achieves functional equivalence to premium commercial RNase R, ensuring complete linear RNA digestion without compromising the integrity of circRNA. The combination of a simplified purification workflow and a robust reaction protocol provides an accessible, cost-effective, and reliable solution for any molecular biology laboratory requiring high volumes of RNase R. Key FeaturesO_LIRNase R mutations can block RNase activity, RNA binding or both C_LIO_LIThis protocol purifies [~]40 mg of active RNase R per liter of E. coli culture C_LIO_LIThe protocol avoids medium and high end FPLC systems C_LIO_LIRNase R expression constructs (WT and mutants) will be available on Addgene C_LIO_LIThe protocol includes an optimized reaction buffer to pair with this RNase R C_LIO_LIOptional endotoxin removal step is also included C_LI

Long-term fiber photometry enables measurement of neural dynamics across hours to days, but these recordings create analytical and reproducibility challenges that are not well addressed by tools developed for short, stimulus-locked experiments. Here we present a software environment for long-term photometry analysis organized around a structured, revisitable workflow for run execution, inspection, and post-run refinement. The software separates correction retuning from downstream event reanalysis, allowing both signal correction and event-analysis settings to be revised after the initial run. We show that correction choice can substantially change the corrected signal itself and that post-run reanalysis can revise event-detection outcomes. The software also preserves tonic and phasic outputs and supports inspection of the same recording at both multiday and session-level scales. Together, these capabilities provide a practical workflow for more interpretable, revisitable, and reproducible analysis of long-term photometry recordings.

The binding of transmembrane (TM) ligands to their cognate TM receptors on neighboring cells governs intercellular adhesion and direct cell-cell communication. However, these interactions are difficult to study in vitro because they depend on membrane presentation, ligand orientation, receptor clustering, and avidity, features often not captured by soluble recombinant ligands or cell-free assays. Here, we describe a flow cytometry-based assay using fluorescent, lentiviral-derived virus-like particles (VLPs) displaying TM ligands to quantify binding to their receptors on target cells. Fluorescent VLPs are generated in-house by plasmid transfection in HEK293T cells and enable direct fluorescent detection without fluorochrome-conjugated secondary antibodies. The system is modular and readily accommodates engineered ligand constructs, including patient-derived variants. We applied this platform to generate ICAM-1-displaying fluorescent VLPs and to study human LFA-1 function in patient-derived leukocytes. This protocol provides a detailed workflow for VLP production and in vitro binding assays, offering a simple, quantitative, and cost-effective approach for studying TM ligand-receptor interactions in a membrane context. The system is well suited for mechanistic studies, functional assessment of patient-derived variants, and direct binding assays using patient-derived cells. Integrating the assay into multicolor flow cytometry panels enables simultaneous immunophenotyping and quantification of up to four ligand-receptor interactions at single-cell resolution. Key featuresO_LIQuantifies TM ligand-receptor binding in a membrane context using fluorescent VLPs and flow cytometry. C_LIO_LIFully in-house, modular system based on plasmid transfection in HEK293T cells, without reliance on recombinant ligands or fluorochrome-conjugated secondary antibodies. C_LIO_LISupports testing of engineered ligand variants, including patient-derived alleles, and direct functional studies on patient-derived cells. C_LIO_LICompatible with multicolor flow cytometry panels, enabling simultaneous immunophenotyping and quantification of up to four ligand-receptor interactions at single-cell resolution. C_LI Graphical overview O_FIG O_LINKSMALLFIG WIDTH=197 HEIGHT=200 SRC="FIGDIR/small/725198v1_ufig1.gif" ALT="Figure 1"> View larger version (55K): org.highwire.dtl.DTLVardef@a43069org.highwire.dtl.DTLVardef@166491borg.highwire.dtl.DTLVardef@49c7d4org.highwire.dtl.DTLVardef@1de36a0_HPS_FORMAT_FIGEXP M_FIG C_FIG

Tracking Single Nucleotide Polymorphisms (SNPs) following CRISPR-Cas9 genome editing is a critical yet often labor-intensive step in modern genetic research. Although Sanger sequencing is the conventional method for definitive confirmation, it typically requires substantial time to generate results. In contrast, PCR-based restriction methods like CAPS (Cleaved Amplified Polymorphic Sequence) and dCAPS (derived CAPS) offer rapid and cost-effective alternatives. However, existing dCAPS primer design tools suffer from significant limitations and were largely developed for tracking polymorphisms in plant genomes. Concurrently, CRISPR-Cas9 gene editing requires strategies to prevent the re-cleavage of the edited allele, typically involving the modification of the Protospacer Adjacent Motif (PAM). To address these challenges, we developed EasyCAPS, a web-based tool that integrates dCAPS primer design with advanced functionalities for CRISPR experiments. EasyCAPS overcomes the shortcomings of previous software by enabling restriction enzyme pre-selection and optimizing designs for complex DNA sequences. Its key innovation is the "Hiding PAM" feature, which designs synonymous mutations to mask the Cas9 recognition site while accounting for codon usage bias, thereby facilitating one-step allelic exchange. The utility of the tool was demonstrated through practical applications targeting the HTA1, PHO84, and CAT5 genes, significantly accelerating both genotyping and gene editing processes. We conclude that EasyCAPS is an accessible solution that effectively streamlines molecular biology workflows.

MOTIVATIONAdult cardiomyocytes are difficult to profile by whole-cell single-cell RNA sequencing because of their large size and fragility, which make them poorly compatible with standard workflows. Current approaches for adult cardiomyocyte transcriptomics often require a trade-off between data quality and throughput, thus, studies instead rely heavily on sequencing of nuclei alone. Therefore, we set out to develop a high-quality and scalable workflow for adult heart cells using in-cell ligation and split-pool barcoding strategies to address this methodological gap. This workflow may be further generalisable to other large cell types or samples containing cell populations with highly unequal RNA content. SUMMARYAdult cardiomyocytes are difficult to profile by whole-cell single-cell RNA sequencing (scRNA-seq). Here, we developed a high-quality and scalable workflow for adult heart cells using in-cell ligation and split-pool barcoding. We identified per-cell RNA content as a significant variable that must be accounted for. Separation of cardiomyocytes (large cells) and non-cardiomyocytes (small cells) before library construction, and allocation of deeper sequencing to cardiomyocytes, produced high-quality whole-cell datasets for both compartments. Compared with single-nucleus RNA sequencing, whole-cell cardiomyocyte profiling better recovered metabolic, mitochondrial, cytoplasmic translational, and contractile gene programs. This workflow provides a practical method for scalable, high-quality cardiomyocyte whole-cell scRNA-seq and offers general strategies for other large cell types or samples containing cell populations with highly unequal RNA content.

How gene expression patterns change spatially as the embryo transitions from simple to complex structures remains a major developmental biology question. Recently developed imaging-based spatial transcriptomics (ST) enable mapping expression of multiple gene at a single-cell resolution. Although Xenopus is a key model in embryology there is no established ST pipeline, and commercially available techniques face many challenges (sample preparation, probe design, cell segmentation). Furthermore, the highly diverse cell shapes and sizes across developmental stages and between different tissues represent major hurdles to accurately defining cells. Here, we describe an optimized workflow for ST in blastula-to-tailbud-stage frog embryos using Merscope, commercial MERFISH (Multiplexed Error-Robust Fluorescence In Situ Hybridization) originally designed for standard mammalian tissues. With stringent quality control and tailored computational pipelines, we optimize this technology for robust, semi-quantitative profiling of spatial transcriptomic landscapes in non-mammalian embryos. Reliable tissue preservation and cell-segmentation enable high-resolution mapping of gene expression during the development of a complex multi-tissue organization. This versatile strategy applies broadly to various dynamic systems, from embryos of various model organisms to complex and heterogeneous organs in mammals. Summary statementThis Single-cell Spatial Transcriptomics pipeline and reference atlas in Xenopus - a model organism in embryology - overcome technical challenges and resolve dynamic changes in patterning during development.

In vitro cytotoxicity assessments frequently rely on staining-based methods that indirectly estimate viable cell numbers indirectly. 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 to 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 streamlined, universal protocol for implementing the resazurin reduction assay across diverse experimental models, emphasizing its practicality, reproducibility, and adaptability for real-time viability monitoring. Key featuresO_LIReal-time, non-destructive monitoring: Enables longitudinal studies by allowing repeated measurements of the same samples over hours without toxicity or disruption. C_LIO_LIStreamlined workflow: A simple "add-incubate-read" protocol eliminates the need for cell lysis, washing, or extraction, saving time and reducing variability. C_LIO_LIBroad sample compatibility: Versatile and reliable for use with 2D monolayers, 3D spheroids, organoids, and bacterial cultures. C_LIO_LIHigh sensitivity: Fluorescent detection of resorufin provides exceptional sensitivity, enabling accurate quantification of even small viable cell populations. C_LIO_LILow background and minimal interference: A clean fluorescent readout reduces the risk of signal artifacts, offering a more reliable alternative to traditional colorimetric assays. C_LIO_LICost-effective and accessible: Utilizes standard laboratory plate readers and commercially available reagents, making it an economical choice for any lab. C_LIO_LIScalable for high-throughput screening: Easily adaptable to various plate formats, supporting both small-scale experiments and large-scale automated screening applications. C_LI Graphical overview O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=141 SRC="FIGDIR/small/718248v1_ufig1.gif" ALT="Figure 1"> View larger version (56K): org.highwire.dtl.DTLVardef@82bcecorg.highwire.dtl.DTLVardef@14164aforg.highwire.dtl.DTLVardef@395118org.highwire.dtl.DTLVardef@fb1349_HPS_FORMAT_FIGEXP M_FIG C_FIG

Osteocytes play a central role in bone remodeling, mineral metabolism, and skeletal homeostasis, but direct molecular analysis of human osteocytes remains technically challenging because they are embedded within the mineralized bone matrix. Surgically obtained human bone specimens provide valuable material for studying human bone biology; however, surface-associated cells, marrow-derived cells, and adherent soft tissues can confound downstream transcript analysis. Here, we describe a bone fragment-based protocol for preparing surgically obtained human bone specimens for molecular analysis of osteocyte-associated transcripts. The protocol consists of mechanical trimming, mincing into small bone fragments, repeated washing, and five sequential rounds of collagenase digestion to reduce non-osteocytic cellular components associated with the bone surface and marrow spaces. The remaining mineralized bone fragments are then frozen in liquid nitrogen, cryogenically pulverized, and lysed in TRIzol reagent for total RNA extraction. Histological validation using residual maxillary bone specimens showed that sequential collagenase digestion markedly reduced adherent soft tissue and extra-matrix nuclei while preserving osteocyte lacunar occupancy. This protocol provides a practical workflow for bone fragment-based RNA analysis focused on osteocyte-associated transcripts in human bone specimens. Specifications table O_TBL View this table: org.highwire.dtl.DTLVardef@1cec618org.highwire.dtl.DTLVardef@2f746forg.highwire.dtl.DTLVardef@1854247org.highwire.dtl.DTLVardef@1c26c1aorg.highwire.dtl.DTLVardef@1473a88_HPS_FORMAT_FIGEXP M_TBL C_TBL

Neural rosettes are hallmarks of the neural progenitor cell stage that is a necessary pre-condition for manufacturing central nervous system lineages. Characterization of early changes during differentiation through positional arrangement and metabolic shifts that occur in a multi-day protocol would facilitate cell culture quality monitoring and optimization of batch culture yield. We describe an analytical framework for identifying neural rosettes from confocal microscopy within a colony of differentiating stem cells and translating co-registered, cell-resolved MALDI imaging data into interpretable readouts that are compatible with cell manufacturing needs. Rather than evaluating hundreds of ion images sequentially, the pipeline converts each region of interest into a single-cell feature matrix and summarizes whole-spectrum variation using PCA, graph-based Leiden clustering, and UMAP visualization. The resultant metabolic neighborhoods provide quantification of molecular heterogeneity within colonies and - when mapped back to x-y space - form coherent spatial domains. Together, these outputs create a practical bridge between multimodal MALDI capabilities and process-relevant interpretation: neighborhoods can be compared across conditions, ranked markers can be prioritized as putative critical quality attributes, and spatial organization can be quantified without manual, feature-by-feature screening.

DNA-RNA hybrids (R-loops) form transiently on the genome and regulate cellular homeostasis. They also influence genome editing outcomes, highlighting their therapeutic potential in vivo. This protocol enables high-resolution mapping of DNA-RNA hybrids directly from frozen mouse tissues. Following tissue homogenisation and lysis, genomic DNA is extracted, digested and DNA-RNA hybrids are isolated using the hybrid-specific S9.6 monoclonal antibody. The purified hybrids are then processed for whole-genome sequencing to generate R-loop profiles. For complete details on the use and execution of this protocol, please refer to Puzzo, Crossley et al1. O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=188 SRC="FIGDIR/small/716701v1_ufig1.gif" ALT="Figure 1"> View larger version (39K): org.highwire.dtl.DTLVardef@3d8f86org.highwire.dtl.DTLVardef@199cd84org.highwire.dtl.DTLVardef@83c51eorg.highwire.dtl.DTLVardef@1024332_HPS_FORMAT_FIGEXP M_FIG O_FLOATNOgraphical abstractC_FLOATNO C_FIG

FSFC is an emerging technology that can greatly enhance our understanding of the single-cell proteomic landscape. However, its application to cells derived from solid tissues has been hampered by their complex autofluorescence signatures and lack of optimized tools for non-immune cells. Here, we present a protocol and discuss key controls that minimize the impact of unmixing errors enabling us to resolve multiple EC subpopulations isolated from different tissues in models of chronic tissue injury. Research Topic(s)Vascular biology, cell heterogeneity, full spectrum flow cytometry Graphical abstract O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=107 SRC="FIGDIR/small/695385v2_ufig1.gif" ALT="Figure 1000"> View larger version (43K): org.highwire.dtl.DTLVardef@1745181org.highwire.dtl.DTLVardef@1930db9org.highwire.dtl.DTLVardef@16a0b3dorg.highwire.dtl.DTLVardef@107ec29_HPS_FORMAT_FIGEXP M_FIG C_FIG HighlightsOptimisation of a FSFC panel to enable in-depth phenotyping of tissue- and model-specific endothelial subpopulations from solid tissues. Discussion of appropriate controls to minimize the impact of tissue autofluorescence and enhance the signal-to-noise ratio for cell phenotyping in complex models of inflammation and fibrosis. Trajectory analysis to track cellular plasticity over time. Application of full spectrum cell sorting to isolate rare endothelial subpopulations with complex phenotypes.

Proprioception and reflexive control of muscle tone depend on the activity of muscle spindles, specialized sensory receptors embedded deep within skeletal muscle that detect changes in muscle length. Their location and complex three-dimensional architecture have historically limited morphological analysis to techniques such as silver-impregnation, muscle teasing, or serial sectioning followed by volumetric reconstruction. Here, we describe a workflow for three-dimensional, in situ visualization of muscle spindles in the rabbit tenuissimus muscle, a preparation uniquely enriched in spindles and well suited for whole-mount imaging. The protocol combines fluorescent labeling of spindle sensory and motor innervation, including intrafusal {gamma} neuromuscular junctions labeled with -bungarotoxin, with immunolabeling and solvent-based optical clearing. Optically cleared tenuissimus muscles were compatible with both whole-mount confocal and light-sheet microscopy, enabling volumetric imaging of complete spindle structures and detailed visualization of Ia annulospiral endings at the spindle equator. This approach provides access to spindle morphology and connectivity at multiple spatial scales while avoiding physical sectioning and reconstruction. By enabling reproducible three-dimensional imaging of intact muscle spindles, this workflow offers a practical platform for studying spindle structure and plasticity in health and disease.

Accurate verification of transgenic plant materials is essential for maintaining scientific integrity and ensuring experimental reproducibility. As the number and diversity of transgenic constructs continue to expand, there is a growing need for practical and scalable methods that enable routine confirmation of transgene presence and identity. Reliable detection systems are particularly important for laboratories handling large numbers of genetically modified lines or distributing materials across research groups. To address this need, we developed two complementary methods for efficient detection of commonly used transgenes. The first method, fDET, is a higher-throughput system capable of simultaneously detecting 15 transgenes and three endogenous genes in a single multiplex PCR reaction followed by capillary electrophoresis. This approach provides rapid, high-resolution detection suitable for high-volume or time-sensitive applications. The second method, DET, offers a more accessible workflow that detects 10 transgenes and one endogenous gene using four multiplex PCR reactions followed by agarose gel electrophoresis. Because DET requires only standard molecular biology equipment, it can be readily implemented in a wide range of laboratory environments without specialized instrumentation. Together, these methods provide flexible and practical solutions for verifying the genetic status of both transgenic and non-transgenic plant materials. By enabling efficient and comprehensive transgene detection, they support reproducible experimentation, facilitate quality control in plant research, and streamline the management and exchange of genetically modified lines. These approaches contribute to more reliable and transparent use of transgenic resources across the plant science community.

Currently there is a lack of high-throughput, low material-input methods to screen early-stage product quality of viral and non-viral gene therapy products. Here we propose using multiplex droplet digital PCR (dPCR) to screen and characterize vector sequences. We describe the adaptation of a Poisson-multinomial model to quantitate integrity of any combination of 4 targets in multiplexed ddPCR. We show the success and limitations of model employment and provide some suggested best practices.

Investigating single-cell dynamics and morphology in tissues and embryos requires highly accurate quantitative analysis of microscopy images. Despite significant advances in the field of bioimage analysis, even the most sophisticated segmentation and tracking algorithms inevitably produce errors (e.g. : over segmentation, missing objects, miss-connected objects). Although error rate may be small, their propagation throughout a time-lapse sequence has catastrophic effects on the accuracy of tracking and extraction of single cell parameters. Extracting single cell temporal information in the context of tissue/embryo requires thus expert curation to identify and correct segmentation errors. In the movies commonly used in developmental biology and stem cell research, both the number of imaged cells and the duration of recording are large, making this manual correction task extremely time-consuming. This has now become a major bottleneck in the fields of development, stem cell biology and bioimage analysis. We present here EpiCure (Epithelial Curation), a versatile tool designed to streamline and accelerate manual curation of segmentation and tracking in 2D movies of large epithelial tissues. EpiCure uses temporal information and morphometric parameters to automatically identify segmentation and tracking errors and provides user-friendly tools to correct them. It focuses on ergonomics and offers several visualization options to help navigating in movies of tissue covering a large number of cells, speeding up the detection of errors and their curation. EpiCure is highly interoperable and supports input from a wide range of segmentation tools. It also includes multiple export filters, enabling seamless integration with downstream analysis pipelines. In this paper, using movies from several animal models, we highlight the importance of curating cell segmentation and tracking for accurate downstream analysis, and demonstrate how EpiCure helps the curation process for extracting accurate single cell dynamics and cellular events detection, making it faster and amenable on large dataset.