STAR Protocols

The bacterial three-hybrid (B3H) assay is a powerful genetic tool for detecting interactions between RNA and RNA-binding proteins (RBPs) and assessing the consequences of RBP mutations. This transcription-based system connects the strength of an RNA-protein interaction to the expression of a lacZ reporter gene in Escherichia coli cells. This in vivo approach allows researchers to dissect RNA-protein interactions within a cellular environment, bypassing the need for biochemical purification of RNAs or proteins. This chapter details a three-day protocol for generating quantitative B3H data. Since a significant challenge in B3H assays is RNA misfolding, we describe a recently optimized set of B3H constructs that mitigates this issue by isolating bait RNAs as discrete folding units.

This step-by-step protocol provides a cross-species, non-surgical approach that enables prenatal gene delivery to the developing nervous system in rats and marmosets. Under transabdominal ultrasound guidance, intracerebroventricular injection of recombinant adeno-associated virus vectors into the fetal brain achieves robust and long-term transduction from prenatal stages into adulthood. This approach can be adapted to other species and target sites outside nervous system, enabling safe and selective intrauterine manipulation and the generation of diverse experimental models for basic and preclinical research. For complete details on the use and execution of this protocol, please refer to Ribeiro Gomes et al (2026)1. Graphical abstract O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=181 SRC="FIGDIR/small/737050v1_ufig1.gif" ALT="Figure 1"> View larger version (47K): org.highwire.dtl.DTLVardef@696364org.highwire.dtl.DTLVardef@fc3c7forg.highwire.dtl.DTLVardef@1e7c7caorg.highwire.dtl.DTLVardef@1edcef0_HPS_FORMAT_FIGEXP M_FIG C_FIG Before you beginExperimental procedures during gestation allow researchers to study developmental processes, including how manipulations of the fetus and its intrauterine environment influence biological outcomes. Ultrasound imaging guidance greatly facilitates such interventions by providing safe and targeted access to fetal compartments, including for prenatal gene delivery to developing neural cell populations. Critically, delivery of recombinant adeno-associated viruses (rAAVs) into the cerebrospinal fluid (CSF) of developing animals enables widespread gene transfer across the brain. The efficiency and distribution of transduction are strongly influenced by developmental stage, making the timing of delivery an important experimental variable. In altricial species such as mice, major developmental processes, including cortical lamination and the establishment of long-range connections, begin prenatally but continue throughout early postnatal life. In primates, however, development is more advanced at birth, and many equivalent developmental events are shifted to the prenatal period. Consequently, developmental stages that can be targeted postnatally in mice require prenatal access in primates. Here, we present a step-by-step protocol for ultrasound-guided fetal intracerebroventricular viral injection (FIVI) of rAAV in marmosets (Callithrix jacchus) and rats (Rattus norvegicus). The procedure was initially developed and optimized in rats before being translated to marmosets, small New World primates that share key developmental, anatomical, and functional characteristics with humans. Together, these models illustrate the cross-species applicability of the approach, while providing gene delivery strategies for both a genetically tractable rodent model and a translationally relevant nonhuman primate. FIVI enables broad gene transfer and stable, long-term transgene expression in wild type animals, facilitating the generation of complementary quasi-transgenic models for research and translational applications from prenatal development through adulthood.

While previous in vitro porcine (Sus scrofa) oviduct spheroid models often rely on manual selection of the formed aggregates, here, we establish a scalable and reproducible system for generating oviduct spheroids. We describe steps for isolating primary secretory and ciliated oviduct epithelial cells and their assembling them into spheroids by forced aggregation using the AggreWell platform. This enables the mass production of uniformly sized spheroids ready for use in a range of downstream applications, such as sperm-oviduct co-incubation experiments. For complete details on the use and execution of this protocol, please refer to Molina et al.1 Graphical abstract O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=197 SRC="FIGDIR/small/731731v1_ufig1.gif" ALT="Figure 1"> View larger version (35K): org.highwire.dtl.DTLVardef@20f94aorg.highwire.dtl.DTLVardef@ce80eeorg.highwire.dtl.DTLVardef@172c631org.highwire.dtl.DTLVardef@31f420_HPS_FORMAT_FIGEXP M_FIG C_FIG Before you beginThis protocol describes the isolation of primary porcine oviduct epithelial cells and their use in generating a three-dimensional (3D) spheroid model in vitro. The protocol is specifically designed to enrich and preserve ciliated oviduct epithelial cells, which are critical for physiologically relevant sperm-oviduct interaction studies yet are frequently underrepresented in conventional in vitro oviduct models. To minimize the de-differentiation of ciliated cells during culture, a specialized Incubation Medium is employed to maintain epithelial cell viability and ciliary function. The relatively brief spheroid formation period (48 h) further supports preservation of ciliary structure and activity prior to downstream functional applications. This protocol consists of four major steps. Steps 1 and 2 cover oviductal epithelial cell isolation through mechanical and enzymatic digestion, step 3 describes the spheroid formation and culture conditions, and step 4 applies the spheroid model to sperm co-incubations or immunofluorescent staining. Although optimized for the porcine isthmus, this protocol may also be suitable for the cells from the ampulla and adaptable to other large animal species.

The middle cerebral artery (MCA) is critical for cerebral blood flow autoregulation and a primary site of cerebrovascular pathology in stroke, Alzheimers disease, and vascular dementia. Pressure myography enables precise ex vivo quantification of MCA structure and function, but requires accurate anatomical identification and careful vessel handling to ensure reproducibility across diverse rat genetic models. This chapter provides a comprehensive, step-by-step protocol for isolating and cannulating the rat MCA M2 segment for pressure myography. We detail precise anatomical landmarks to ensure consistent vessel selection across strains. The protocol includes optimized solutions, cannulation techniques, and pressure protocols validated across multiple rat models, including transgenic (TgF344-AD), diabetic (T2DN), consomic (SS.5BN, FHH.1BN), and genome-edited strains. Extensive troubleshooting notes address common technical challenges, including vessel viability assessment, pressure integrity, and strain-specific autoregulatory ranges. This methodology bridges molecular genetic findings with fundamental cerebrovascular physiology, enabling researchers to characterize myogenic reactivity, passive mechanical properties, and structural remodeling in rat models of cerebrovascular disease.

CD8 T cells play a central role in immune responses to infection and cancer. However, the diversity of T cell receptor (TCR) specificities makes it challenging to study the mechanisms that regulate T cell activation, differentiation, and effector function. Beyond TCR transgenic mouse models, various complex genome-editing approaches have been employed to overcome this challenge. However, these strategies are often technically demanding, time-intensive, and difficult to adapt. Investigators who are interested in testing de novo TCRs under their chosen experimental conditions would benefit from a standardized and accessible method. Here, we describe a protocol that combines ribonucleoprotein (RNP)-based CRISPR-Cas9 editing with retroviral transduction to enable efficient genetic manipulation of murine CD8 T cells. We show that T cells engineered via this protocol can be generated at sufficient scale for downstream in vitro assays and in vivo adoptive transfer experiments. We expect this method will be useful for investigators who require a standardized and accessible way to study how TCR specificity impacts CD8 T cell responses.

Manual analysis of skeletal muscle cross-sections is time-consuming and subject to error and user bias. To overcome these limitations, we developed and validated a semi-automated, quantitative, and reproducible image-analysis pipeline specifically tailored to quantify Pax7+ satellite cells, myonuclei, and cross-sectional area by fiber type. The workflow combines FIJI/ImageJ-based image preprocessing with CellProfiler, Cellpose, and a custom Python script to process and analyze immunohistological images of muscle tissue cross-sections. Outcomes include Pax7+ satellite cells and myonuclei quantified per fiber by fiber type, along with cross-sectional area, perimeter, and fiber type classification. This semi-automated approach provides a robust and efficient platform for high-throughput analysis of muscle tissue cross-sections from large datasets. O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=141 SRC="FIGDIR/small/729866v1_ufig1.gif" ALT="Figure 1"> View larger version (44K): org.highwire.dtl.DTLVardef@a3401dorg.highwire.dtl.DTLVardef@1c63145org.highwire.dtl.DTLVardef@ccbf76org.highwire.dtl.DTLVardef@2e0da0_HPS_FORMAT_FIGEXP M_FIG C_FIG

Single-cell and single-nucleus RNA sequencing have become increasingly widespread, creating a significant demand for accessible analysis tools in research laboratories. Despite this need, the bioinformatics expertise required for such analyses remains rare. Cell-Hub addresses this gap by enabling single-cell data analysis for all researchers, regardless of computational background. Cell-Hub is a comprehensive, free, and open-source framework built on R/Shiny and distributed as a Docker image, integrating Seurat 5, CellChat 2, and Monocle 3 within a unified graphical interface. It supports all essential steps of single-cell RNA-seq analysis: data loading, quality control, normalization, clustering, multi-dataset integration, differential expression, and biomarker detection. Cell-Hub further incorporates ligand-receptor interaction inference powered by GaspouDB, a consolidated database of 11,563 mouse and 9,604 human interactions derived from CellChat, CellPhoneDB, CellTalkDB, and MultiNicheNet as well as trajectory inference via Monocle 3 and spatial transcriptomics analysis for 10X Visium datasets. All analyses produce publication-ready visualizations with flexible export options. By integrating these analytical frameworks into a single, intuitive interface requiring no programming expertise, Cell-Hub represents a significant step toward democratizing single-cell genomics for the broader research community.

Whole-genome sequencing identifies focal DNA amplifications with base-pair resolution but cannot determine whether amplified sequences reside on extrachromosomal DNA (ecDNA, also known as double minutes) or within chromosomally integrated homogeneously staining regions (HSRs). DNA fluorescence in situ hybridization (DNA-FISH) metaphase spreads remain the gold standard for distinguishing these amplification states at single-cell resolution. Here, we present a detailed protocol for DNA-FISH metaphase spreads using human cancer cell lines, encompassing cell culture, metaphase arrest, hypotonic treatment, fixation, chromosome spreading, fluorescent probe hybridization, and fluorescence imaging. The protocol incorporates intermediate quality-control steps to verify successful chromosome dispersion and optimize metaphase spread quality, making the workflow accessible to laboratories without specialized cytogenetics expertise. Results demonstrate clear visualization of ecDNA and HSR amplification states using locus-specific probes and illustrate common technical artifacts that can affect interpretation. This protocol provides a robust and reproducible approach for studying the structural organization of oncogene amplification in cancer cells.

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 successful application across different cell line spheroids, with subsequent compatibility with hematoxylin and eosin (H&E) staining protocols. Compared to conventional methods, our approach significantly reduces sample loss, improves 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 and molecular assessment in translational research settings. Key featuresO_LIMaintains spheroid integrity: Prevents mechanical distortion, fragmentation, and loss of spatial orientation during processing. C_LIO_LISignificantly reduces sample loss: Decreases failure rate compared to traditional methods, conserving valuable samples. C_LIO_LIBroad spheroid compatibility: Works effectively with primary tumor-derived, stem cell-derived, and co-culture spheroid models. C_LIO_LIEnables high-quality sectioning and staining: Delivers consistent, reproducible sections that are fully compatible with H&E, IHC, and IF. C_LI Graphical overview O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=140 SRC="FIGDIR/small/743094v1_ufig1.gif" ALT="Figure 1"> View larger version (44K): org.highwire.dtl.DTLVardef@1670c4org.highwire.dtl.DTLVardef@145810aorg.highwire.dtl.DTLVardef@1accb1org.highwire.dtl.DTLVardef@17481c0_HPS_FORMAT_FIGEXP M_FIG C_FIG

Native polyacrylamide gel electrophoresis is the standard first analytical step after detergent solubilization of thylakoid membranes. It separates photosystem supercomplexes by size and shape, but it forces a polydisperse mixture of protein-detergent particles through a gel of limited pore diameter. Interval-zone free-flow electrophoresis (iZE-FFE) performs that first electrophoretic step in free solution, without ampholytes or immobilines, and returns the complexes as a liquid fraction series ordered by net charge. We describe the operational cycle of iZE, show that bromophenol blue reports the charge density of {beta}-dodecylmaltoside micelles, and apply iZE to Arabidopsis thaliana thylakoid extracts from two sequential solubilization campaigns. After a two-step digitonin then digitonin/{beta}-dodecylmaltoside extraction (experiment 6874), SDS-PAGE of the iZE fractions is crowded: high-molecular mass protein-lipid assemblies of photosystem I and photosystem II release the same subunits into many wells. Native-PAGE of the same fractions resolves ATP-synthase from PSI, a free LHCII pool, anodic PSII, and cathodic PSI, including PSI with and without LHCII. A three-step extraction with a single pH working window (experiment 7154) confirms this charge order. Lowering residual detergent in consecutive extracts still produces partly solubilized high-mass assemblies, whereas a final {beta}-dodecylmaltoside step releases the C2S2M2/C2S2M/C2S2 PSII series and a single PSI band. iZE is therefore a charge-first, orthogonal dimension for native thylakoid biochemistry. Native-PAGE, not SDS-PAGE, is the proper second dimension while solubilization remains incomplete.

Click-qPCR is a browser-based application for relative qPCR analysis that requires a tidy-format CSV file containing four columns: sample, group, gene, and Cq. Preparing this input from qPCR instrument output typically requires manual reformatting and calculation of mean Cq values for technical replicates. To simplify this process, we developed Click-Prep (https://kubo-azu.shinyapps.io/Click-Prep/), an interactive web-based application designed specifically to create Click-qPCR input files. Click-Prep imports CSV, TXT, TSV, and XLS/XLSX files and supports skipping of instrument-generated metadata rows, interactive column mapping, and manual assignment of experimental groups. Users can review technical-replicate measurements, exclude selected rows according to predefined quality-control criteria, and calculate mean Cq values for each sample-group-target combination. Missing or nonnumeric Cq values are flagged for review and must be resolved before the mean is calculated. Click-Prep can also combine compatible formatted CSV files, such as datasets obtained from separate qPCR plates. The resulting dataset is exported as a standardized CSV file containing the four fields required by Click-qPCR. By integrating these operations into a guided browser-based workflow, Click-Prep enables users to prepare Click-qPCR input files rapidly and consistently without programming.

We describe here an efficient method for gene editing in mouse T cells, based on well-established, high-efficiency retroviral transduction protocols. Our platform allows analysis of mutant phenotypes in primary murine T cells in vitro and in vivo. This approach uses a single retroviral vector to simultaneously knockout an endogenous gene and ectopically express its mutant version. This knockout/re-expression vector can be used as the only plasmid to transduce Cas9-expressing T cells, or used together with a Cas9 retroviral vector to transduce T cells from any mouse strain. We validated the system for analysis of murine T cells by targeting key molecules in proximal T cell signaling, i.e. CD3{gamma} and Zap70. We obtain high knockout and re-expression efficiencies in both Cas9-expressing and non-Cas9 T cells. Knockout efficiencies can be further improved by gRNA multiplexing. Endogenous proteins compete with their ectopically expressed mutants or tagged versions for cellular location, protein interactions and cellular functions. Here, we quantified the incorporation of CD3{gamma}-GFP into surface T cell receptor (TCR) complexes. Our data shows that the knockout and re-expression platform improves integration of CD3{gamma}-GFP into the TCR. Therefore, eliminating competition between endogenous and ectopic proteins benefits analyses of protein assemblies and signaling pathways in primary T cells. Furthermore, we validated advantages of our system for mutant analysis using wild-type and mutant Zap70s. Zap70 mutants deficient in TCR binding or kinase activity show their phenotypes only in the absence of endogenous protein, further validating our knockout/re-expression approach. Most importantly, this system can be used to generate gene-edited primary T cells for in vivo studies, such as the quantification of anti-tumor responses. Our knockout and re-expression platform provides a useful gene editing tool for primary T cells in fundamental research and immunotherapy development.

Derived Cleaved Amplified Polymorphic Sequences (dCAPS) assays have been widely performed historically to detect known base substitutions in many model organisms--notably Caenorhabditis elegans, Arabidopsis thaliana, Saccharomyces cerevisiae, and Schizosaccharomyces pombe-- where chemical mutagens that induce point mutations are frequently used. With the rise of whole-genome sequencing and genome editing technologies, dCAPS is increasingly applied to detect diverse nucleotide changes in additional organisms, including Drosophila, zebrafish, mammals, and agricultural crops (e.g., Oryza sativa and Hordeum vulgare). However, a key limitation of dCAPS is that genomic target sites amenable to primer designs that both preserve PCR amplification and create recognition sites for inexpensive, high-performance restriction enzymes are scarce. Here we report One-step dual-primer dCAPS (OddCAPS), a modification that uses three primers in one reaction to overcome this constraint. Two of these primers, an intermediate primer and a dCAPS primer, sequentially introduce 1-2 base substitutions each into the amplicon, enabling up to four engineered base changes near the nucleotide of interest. By using the intermediate primer at 1/10-1/100 the concentration of the other primers, the desired product is generated directly in a single-tube, one-step PCR. Increasing the number of engineered substitutions improves the chance of using a researchers preferred restriction enzyme. In principle, having eight common restriction enzymes (BamHI, EcoRI, NheI, SalI, BglII, ClaI, HindIII, and MluI) suffices to detect any single-nucleotide variant in any biological or synthetic DNA sequence with this approach.

Single-cell RNA sequencing now routinely produces detailed maps of cell types and states, but interpreting a finished project remains harder than it should be. Once the analysis is done, the results are usually handed over as static reports, figure panels and supplementary tables. A biologist who later wants to revisit an annotation, recompute a cell-type proportion or check whether a pathway is specific to one group typically has to return to a bioinformatician rather than explore the data directly. We developed scSpark to close this gap. The platform takes the completed outputs of a single-cell project: cell annotations, embeddings, differential-expression tables, trajectories, cell-cell communication networks and enrichment result and serves them through a web browser as an interactive workspace. Heavy computation stays upstream: scSpark indexes the precomputed objects under a single project structure and exposes them through six modules for cell annotation, differential analysis, trajectory exploration, cell-cell communication, functional interpretation and AI-assisted result interrogation. Every action in these modules, from a query to a label change, an export or an AI-generated summary, is linked to a specific project version, data object, parameter set and output file, so that any conclusion can be traced back to the evidence behind it. We illustrate the platform by reworking a published periodontitis dataset through this interface. scSpark does not replace upstream pipelines or expert judgement; it is a layer that makes their results easier to inspect, revise and reuse, and that turns a single-cell project from a one-off report into an interpretation others can follow and check.

Here we introduce OpenEvo, a fully open-source, low-cost turbidostat platform for automated continuous culture and directed evolution experiments. Existing tools are expensive, complex, or lack open-source hardware; OpenEvo addresses this gap. OpenEvo is a complete, fully automated evolution platform with detailed, illustrated construction instructions for beginners, open-source software and firmware, and a single device priced around $300. An optional PC-based version offers enhanced functionality, including remote access, programmable evolution cycles, programmable LED stimulation, and a data visualization tool. OpenEvo can cycle through three types of media for positive, negative, and neutral selection conditions, supporting a wide range of experimental designs. We validate the use of OpenEvo by evolving H. volcanii to grow from 15% to 12% salt over ~150 cycles, ~1,000 hours. Evolved cells grew 36% faster than wild-type at 12% salt. Whole-genome sequencing of adapted cells found SNPs and large deletions. We also demonstrate positive and negative selection using the OpenEvo LEDs to drive optogenetics via a Phytochrome B-based optogenetic tool, with light as the selection stimulus during over 4000 hours of growth. OpenEvo lowers the technical and cost barriers for continuous evolution experiments, serves as a teaching tool, and is designed to grow an open community of users who share modifications.

GapAutomated spike sorting algorithms have revolutionized the way neuronal activity is extracted from extracellular recordings, yet they remain imperfect. Specifically, inaccurate acceptance of noise-based units not only leaves researchers with clusters that require extensive manual curation, an essential but time-consuming process, that also leads to significant subjectivity in the selection of units. In an era of high-density probes like Neuropixels, where an hour of data can exceed 80 GB, manual curation is no longer scalable, automation of standard criteria can speed data curation and ensure quality of datasets. Here, we developed a semi-automated curation pipeline to label the quality of units after automated curation by Kilosort. ApproachOur algorithm standardizes criteria for labeling of Noise, Multi-Unit Activity (MUA), and Good Units using a combination of spike rate, spike timing metrics (from autocorrelogram), and waveform-based physiological features such as peak amplitude, slopes, half-width, and inter-channel correlation. Based on these features, clusters are assigned standardized labels (good, noise, multi-unit activity) that can be imported directly into Phy, where they serve as curation aids rather than absolute classifications, supporting but not replacing expert judgment. Heuristically, "noise" units are those unlikely to be neuronal in origin; "MUA" includes units with significant neural contribution (i.e., neuronal waveform) but with some degree of clear imperfection to be further cleaned, and "good" units are those without any clear deviation from ideal unit criteria. By ensuring accurate selection of acceptable units, we enable robust downstream analyses such as neural decoding and longitudinal tracking of neuron identity. Thresholds for all metrics were chosen to maximize the matching of algorithm output to that of 2 expert manual curators. Of note, users may alter thresholds either based on their own judgment or using an included tool to semi-automatically find thresholds that optimize SpikeCleaner with their own expert curation. Results: To benchmark, we compared the outputs of our algorithm to expert-labels curated in Phy by two expert users across three recordings. SpikeCleaner achieved an average of 97% accuracy vs. experts & 92% F1 score in classifying Single Units. It achieved an accuracy of 97% & 92% F1 score in full-category agreement (SU, MUA, Noise), and 97% accuracy & 95% F1 score in distinguishing Neuronal vs. Non-Neuronal units.

Circular single-stranded DNA (cssDNA) is a versatile biomolecule with applications spanning genome editing, DNA nanotechnology, synthetic biology, molecular diagnostics, and aptamer development. Compared with linear single-stranded DNA, cssDNA offers enhanced structural stability, resistance to in-cellulo degradation by exonucleases and enables the generation of long, sequence-defined DNA molecules that are difficult to obtain through conventional chemical synthesis methods. Despite its growing utility, widespread adoption of cssDNA has been limited by the lack of accessible, scalable, and cost-effective production methods, with many existing workflows relying on specialised reagents, extensive optimisation, or commercially synthesised DNA. Here, we present a streamlined, end-to-end protocol for the laboratory-scale production of high-purity cssDNA using an M13 phagemid-based system and standard molecular biology laboratory infrastructure. The workflow encompasses bacterial culture, phage amplification, nuclease treatment, phage precipitation, anion-exchange purification, and quality control, with practical optimisations to improve yield, reproducibility, and scalability. Using this approach, yields range from 120-195 {micro}g of purified cssDNA from 300 mL of culture supernatant. The protocol provides detailed guidance on critical process parameters, troubleshooting, and quality assessment, enabling reliable production of cssDNA suitable for a wide range of downstream molecular biology and genome engineering applications.

The human small intestine (SI) plays a central role in nutrient processing, host-microbe interactions, and immune regulation, yet remains poorly characterized due to the lack of minimally disruptive sampling methods. Here, we present a protocol for deploying, recovering, and analyzing samples collected using an ingestible device that enables multi-region, lumen-targeted SI sampling during normal digestion. The device incorporates a ~30-cm collapsible tube wound into pH- or time-responsive layers that sequentially unfurl in situ, typically capturing three spatially ordered samples with high yield and reliable retrieval. This protocol outlines study design, participant handling, device recovery, contamination control, and standardized workflows for analyses, including cell quantification, culturomics, sequencing, and metabolomics. We further describe benchmarking approaches for evaluating spatial resolution and strategies for assay prioritization when sample volume is limiting. By reducing participant burden and facilitating integration with stool, saliva, and clinical metadata, this approach enables longitudinal and large-cohort studies linking SI microbial ecology and host physiology to human health.

Antibody- and cell-mediated immune responses against biologics, should they occur, can impact treatment efficacy and potentially pose severe risks to patient safety. Therefore, developers have focused on advancing strategies to mitigate such unwanted immunogenicity. Opportunities to address immunogenicity early in the development process, particularly during the drug design phase, have been identified. In vitro and in silico tools that facilitate the identification and removal of sequence liabilities have been established. For example, human cell-based in vitro T cell assays can be used to identify and remove CD4+ T cell epitopes, which are known to play a critical role in the development of anti-drug antibodies against recombinant proteins products as well as the transgenes of gene and therapy. Despite their widespread use in the industry, most of these assays lack thorough characterization, which undermines confidence in the results and comparability across laboratories. In this study, concepts of immunogenicity bioanalytical assay validation for study design and analysis were applied to characterize an internal CD4+ T cell proliferation assay as fit-for-purpose. A statistical path was applied to establish data acceptance criteria for handling of replicates, positivity and negativity of a signal, and donor cohort size. A Bayesian analysis was also performed and is proposed as an approach for sequence de-risking decision making. The in-depth characterization of the CD4+ T cell proliferation assay described here allows for accurate interpretation of the assay outcomes, thereby enhancing confidence in using this approach for mitigating the immunogenicity of biologics by design.

BackgroundInvestigating the subcellular distribution of proteins is crucial for understanding complex cell behaviours and disease mechanisms, and fluorescence microscopy has become a key tool for visualising protein localisation. Endogenous protein tagging, where the sequence for a tag (typically a peptide or fluorescent protein) is integrated into the native genetic sequence encoding a protein of interest, enables proteins to be visualised without the need for antibodies against the target protein. ORANGE (Open Resource for the Application of Neuronal Genome Editing) is a CRISPR-Cas9-based endogenous protein tagging technique which relies on homology-independent targeted integration (HITI)-mediated gene editing. Utilising HITI as the DNA repair pathway of choice gives ORANGE the advantage of being more efficient than classical homology-directed repair (HDR)-based endogenous protein tagging techniques and additionally, means it can be used in post-mitotic cells. ResultsWe applied the ORANGE system to tag three proteins, CYFIP1, JAKMIP1, and STAT3, and confirmed that the expressed fusion proteins demonstrate expected subcellular localisations through fluorescence microscopy. Unexpectedly, the efficiency of ORANGE editing was less than 1% in HEK293 cells, despite high transfection efficiency. To improve the editing efficiency associated with ORANGE, we combined the ORANGE method with an established Sleeping Beauty transposase/CRISPR-Cas9 fusion technique, which has been shown to enhance HITI-mediated gene editing. Using this new method, which we term Sleeping ORANGE, we successfully tagged CYFIP1 with the fluorescent protein mNeonGreen. Importantly, quantitative analysis by fluorescence microscopy and flow cytometry demonstrated an increase in editing efficiency using Sleeping ORANGE, with an approximately 12.85-fold increase in the percentage of mNeonGreen-expressing cells at 72 hours post-transfection relative to populations of cells edited with the ORANGE method. ConclusionsWe have incorporated the DNA-binding domain of the Sleeping Beauty transposase to create a new system that improves the gene-editing efficiency of the ORANGE technique. With further developments to optimise CRISPR gRNA design and reduce off-target effects, the Sleeping ORANGE technique may form a valuable tool for researchers to better understand subcellular localisation and dynamics.