STAR Protocols

In the accompanying chapter (Gressel, Lidschreiber, Cramer), we describe the detailed experimental protocol for transient transcriptome sequencing (TT-seq). TT-seq detects metabolically labeled, newly synthesized RNA fragments genome-wide in living cells. TT-seq can monitor gene activity and the dynamics of enhancer landscapes with great sensitivity, but this requires careful computational analysis of the data. In this manuscript, we present the bioinformatics workflow used to analyze TT-seq data. In particular, we describe pre-processing steps, including a reliable and robust normalization strategy, and several downstream analysis tools that enable the user to quantify RNA synthesis, splicing and degradation activities. Together, these tools form a comprehensive analysis pipeline that can be adapted to almost any TT-seq application.

Genetic assays are an invaluable tool for both fundamental biological research and translational applications. Variable Dose Analysis (VDA) is an RNAi-based method for cell-based genetic assays that offers several advantages over approaches such as CRISPR and other RNAi-based methods including improved data quality (signal-to-noise ratio) and the ability to study essential genes at sub-lethal knockdown efficiency. Here we report the development of three new variants of the VDA method called high-throughput VDA (htVDA), VDA-plus and pooled-VDA. htVDA requires 10-fold reduced reagent volumes and takes advantage of liquid handling automation to allow higher throughput screens to be performed while maintaining high data quality. VDA-plus is a modified version of VDA that further improves data quality by 4.5-fold compared to standard VDA to allow highly sensitive detection of weak phenotypes. Finally, Pooled VDA allows greatly increased throughput by analysing multiple gene knockdowns in a single population of cells. Together, these new methods enhance the toolbox available for genetic assays, which will prove valuable in both high-and low-throughput applications. In particular, the low noise and ability of VDA to study essential genes at sub-lethal knockdown levels will support identification of novel drug-targets, among which essential genes are often enriched. While these tools have been developed in Drosophila cells, the underlying principles are transferrable to any cell culture system.

The authors wish to withdraw this manuscript and apologize for errors in the initial submission. All the original experiments were performed by YY. Unfortunately, JP and members of the Mello lab have not been able to replicate some aspects of the study. JP has failed to independently reproduce the specific results showing RNAi-triggered relocalization of target RNA, and P granule specific accumulation of (the P granule component GLH-1) as reported. The conditions/strains analyzed by JP were as follows: (1) oma-1 FISH on WT worms [control, 6 hr and 12 hr oma-1(RNAi)]. (2) oma-1 FISH on OMA-1:GFP worms [control, 6 hr oma-1(RNAi), or 6 hr gfp(RNAi)]. (3) oma-1 FISH on WT worms [control, 4hr, 6hr, 8 hr, and 10 hr oma-1(RNAi)]. 10-23 gonads were analyzed per experiment. Fixation conditions were essentially as described, with the only known difference being that gonads were not exposed to detergent prior to fixation. Using YYs reagents and protocol the Mello lab has not observed an obvious relocalization of target RNA to P granules (marked by GFP::GLH-1) after 6 hrs oma-1(RNAi); n=92 gonads. CM, JP and DG consider that the published images accurately represent the image stacks provided by YY as representative, raw data, but JP and CM note configurations of FISH signals in germ nuclei and gonad anatomy that they consider unusual. CM, JP and DG have not detected any evidence of image manipulation. YY states that none of the raw image data were manipulated beyond standard adjustments for brightness and contrast prior to processing images for publication as described. However, YY reports that the images were not representative of the majority of sample gonads, and instead were pre-selected under low magnification for rare examples with asymmetrical, expanded P granules. Efforts to identify conditions that explain the rare gonads imaged by YY continue in the Mello lab, as do efforts to reproduce independently each of the other reported results; we plan to provide an update in the near future.

Visualization of gene products in Caenorhabditis elegans has provided insights into the molecular and biological functions of many novel genes in their native contexts. Single-molecule Fluorescence In Situ Hybridization (smFISH) and Immunofluorescence (IF) visualize the abundance and localization of mRNAs and proteins, respectively, allowing researchers to elucidate the localization, dynamics, and functions of many genes. Here, we describe several improvements and optimizations to existing IF and smFISH approaches specifically for use in C. elegans embryos. We present 1) optimized fixation and permeabilization steps to preserve cellular morphology while maintaining probe and antibody accessibility, 2) a streamlined, in-tube approach that negates freeze-cracking, 3) the smiFISH (single molecule inexpensive FISH) adaptation that reduces cost, 4) an assessment of optimal anti-fade products, and 5) straightforward quantification and data analysis methods. Most importantly, published IF and smFISH protocols have predominantly been mutually exclusive, preventing exploration of relationships between an mRNA and a relevant protein in the same sample. Here, we present methods to combine IF and smFISH protocols in C. elegans embryos including an efficient method harnessing nanobodies. Finally, we discuss tricks and tips to help the reader optimize and troubleshoot individual steps in each protocol.

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@417649org.highwire.dtl.DTLVardef@1bcd186org.highwire.dtl.DTLVardef@15fecb3org.highwire.dtl.DTLVardef@acdf8d_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.

The isolation of specific cell types of the brain is essential to study cell-type-specific differences in complex neurological diseases such as Alzheimers disease. This protocol isolates oligodendrocytes, microglia, endothelial cells, astrocytes, and neurons from a single mouse brain. The process involves gentle tissue homogenization, debris removal, and sequential sorting of five distinct cell types. We validate cell purity and viability using flow cytometry and RT-qPCR. This protocol is well-suited for a range of downstream applications, including genomics, transcriptomics, and proteomics. O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=200 SRC="FIGDIR/small/666877v1_ufig1.gif" ALT="Figure 1"> View larger version (47K): org.highwire.dtl.DTLVardef@1426b97org.highwire.dtl.DTLVardef@1a5bb77org.highwire.dtl.DTLVardef@1b69f32org.highwire.dtl.DTLVardef@8da416_HPS_FORMAT_FIGEXP M_FIG C_FIG

BackgroundGenome editing in human skeletal muscle research requires protocols that maximize delivery while preserving viability and clonal outgrowth. We sought to develop a reagent-free workflow for CRISPR/Cas9 editing in human immortalized myoblasts and to demonstrate its performance in two use cases, an IARS1 knockout and an MLIP homozygous knock-in. MethodsWe optimized electroporation parameters using a green fluorescent protein reporter to compare three electrical settings for transfection and survival in E6/E7 myoblasts, then applied ribonucleoprotein delivery for editing. We evaluated the effect of confluency at electroporation, performed single-cell cloning without antibiotics or fluorescence-activated sorting, and validated edits by high-resolution melting pre-screen followed by Sanger sequencing. ResultsElectroporation optimization identified one parameter set that maximized delivery while preserving viability. Performing electroporation at low confluency increased clonal outgrowth and editing rates. The workflow yielded an 84% success rate for IARS1 knockout and a 3.3% success rate for MLIP homozygous knock-in. High-resolution melting provided a very sensitive pre-screen, detecting 96% to 100% of actual edits, reducing the number of Sanger sequencing needed. Performance was reproducible across runs and myoblast lines and increasing single-cell seeding scaled yields without compromising purity. ConclusionsThis work provides a practical and reproducible selection-free protocol that couples electroporation optimization, low confluency editing, single-cell cloning, and high-resolution melting sorting to generate pure edited myoblast lines. The approach is applicable to disease modeling in neuromuscular research and clarifies feasibility boundaries for essential genes and homology-directed repair in these cells.

The ability to generate intact nuclei is crucial to the success of a variety of genomics experiments, such as Assay for Transposase-Accessible Chromatin using sequencing (ATAC- seq), Cleavage Under Targets and Tagmentation (CUT&Tag), and nuclei-based single cell sequencing (e.g., single nuclei ATAC-seq and single nuclei RNA-seq). For plants, the presence of the cell wall presents significant challenges in the isolation of nuclei from tissues. Here, we report an optimized nuclei isolation protocol that can be adapted for diverse angiosperm species, including maize, soybean, tomato, potato, and wheat, starting from fresh or frozen tissues. Nuclei release is achieved through chopping tissue on ice, where a key parameter affecting nuclei integrity is the concentration of detergent TritonX-100 in the nuclei isolation buffer. The method is simple, quick, and largely centrifugation-free, in which debris is removed by serial filtration. Initial nuclei release and filtration can be performed within 20 min. Fluorescence activated nuclei sorting is then used for final nuclei purification to remove other organelles such as plastids. The protocol uses 500 mg or less plant tissue as input and typically yields at least 100,000 - 200,000 purified nuclei per sample, a common input amount for downstream experiments. Throughout the protocol, we provide guidelines for optimization if performing nuclei isolation from a given species and tissue for the first time.

SummaryImmunofluorescent staining is commonly used to generate images to characterise cytological phenotypes. The manual quantification of DNA double-strand breaks and their repair intermediates during meiosis using image data requires a series of subjective steps, from image selection to the counting of particular events per nucleus. Here we describe synapsis, a Bioconductor package, which includes a set of functions to automate the process of identifying meiotic nuclei and quantifying key double-strand break formation and repair events in a rapid, scalable and reproducible workflow, and compare it to manual user quantification. The software can be extended for other applications in meiosis research, such as incorporating machine learning approaches to categorise meiotic substages. Availability and implementationsynapsis can be freely downloaded and installed at: http://bioconductor.org/packages/release/bioc/html/synapsis.html R functions and further information can be found at https://gitlab.svi.edu.au/drr-public/synapsis Contactwcrismani@svi.edu.au

One obvious feature of life is that it is highly dynamic. The dynamics can be captured by movies that are made by acquiring images at regular time intervals, a method that is also known as timelapse imaging. Looking at movies is a great way to learn more about the dynamics in cells, tissue and organisms. However, science is different from Netflix, in that it aims for a quantitative understanding of the dynamics. The quantification is important for the comparison of dynamics and to study effects of perturbations. Here, we provide detailed processing and analysis methods that we commonly use to analyze and visualize our timelapse imaging data. All methods use freely available open-source software and use example data that is available from an online data repository. The step-by-step guides together with example data allow for fully reproducible workflows that can be modified and adjusted to visualize and quantify other data from timelapse imaging experiments. O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=194 SRC="FIGDIR/small/432684v1_ufig1.gif" ALT="Figure 1"> View larger version (51K): org.highwire.dtl.DTLVardef@d04086org.highwire.dtl.DTLVardef@3c304borg.highwire.dtl.DTLVardef@186bf56org.highwire.dtl.DTLVardef@17bbd6d_HPS_FORMAT_FIGEXP M_FIG C_FIG

In situ hybridization visualizes RNA in cells, but image analysis is complex. We present a protocol based on open-source software for automated high-content multiplex fluorescence in situ transcriptomics analysis. Steps include nuclei segmentation with a Fiji macro and quantification of up to 14 mRNA probes per image. We describe procedures for storing raw data, quality control images and the use of a Python app to summarize all the results in one spreadsheet detailing the number of single or co-positive cells.

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.

Covaris g-TUBEs can be used to fragment DNA to pre-determined sizes based on the relative centrifugal force that they are run at. They are recommended for use while preparing Oxford Nanopore Technology libraries by the manufacturer. However, the volumes and DNA concentration typically used for ONT libraries are outside the range of the example data provided by Covaris. Here, we ran g-TUBEs at three different relative centrifugal forces and determined the effect on DNA fragmentation in the range 0.5 - 4 {micro}g. This dataset can be used to inform the effective fragmentation of DNA for creating Oxford Nanopore libraries of an optimal size.

Single-nucleus RNA sequencing enables high-resolution transcriptomic profiling of brain tissue, facilitating detailed analysis of cell identity in models of neurodegeneration and repair. Here, we describe a protocol for isolating nuclei from long-term human stem cell-derived grafts in the rat brain, incorporating vibratome sectioning, graft dissection, nuclear extraction, and fluorescence-activated sorting. This workflow supports analysis of human neurons embedded within host tissue or sensitive to dissociation, offering a powerful approach to assess graft composition, integration, and neuronal identity in living brain. For complete details on the use and execution of this protocol, please refer to Fiorenzano et al.1 Subject areasSingle nucleus RNA sequencing, stem cell biology, neuroscience, transplantation, cell replacement therapy Graphical abstract O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=61 SRC="FIGDIR/small/672369v1_ufig1.gif" ALT="Figure 1"> View larger version (18K): org.highwire.dtl.DTLVardef@1ac3b90org.highwire.dtl.DTLVardef@7aab65org.highwire.dtl.DTLVardef@18a8fd4org.highwire.dtl.DTLVardef@1e8bfae_HPS_FORMAT_FIGEXP M_FIG C_FIG HighlightsO_LIStep-by-step vibratome sectioning of xenografted rat brain tissue. C_LIO_LIPrecise dissection of human stem cell-derived grafts from host brain sections. C_LIO_LIExtraction of intact nuclei from fragile, grafted neurons. C_LIO_LIIsolation of single nuclei via fluorescence-activated nuclei sorting (FANS) for snRNA-seq sample preparation. C_LI

Multiparametric imaging allows researchers to measure the expression of many biomarkers simultaneously, allowing detailed characterization of cell microenvironments. One such technique, CODEX, allows fluorescence imaging of >30 proteins in a single tissue section. In the commercial CODEX system, primary antibodies are conjugated to DNA barcodes. This modification can result in antibody dysfunction, and development of a custom antibody panel can be very costly and time consuming as trial and error of modified antibodies proceeds. To address these challenges, we developed novel tyramide-conjugated DNA barcodes that can be used with primary antibodies via peroxidase-conjugated secondary antibodies. This approach results in signal amplification and imaging without the need to conjugate primary antibodies. When combined with commercially available barcode-conjugated primary antibodies, we can very quickly develop working antibody panels. We also present methods to perform antibody staining using a commercially available automated tissue stainer and in situ hybridization imaging on a CODEX platform.

An effective method for tissue specific ablation in zebrafish is the nitroreductase/metronidazole system. Expressing bacterial nitroreductase (ntr) in the presence of nitroimidazole compounds causes apoptotic cell death, which can be useful for understanding many biological processes. However, this requires tissue specific expression of the ntr enzyme, and many tissues have yet to be targeted with transgenic lines that express ntr. We generated a transgenic zebrafish line expressing ntr in differentiated skeletal muscle. Treatment of embryos with metronidazole caused muscle specific cell ablation. We demonstrate this line can be used to monitor muscle regeneration in whole embryos and in transplanted transgenic cells. Summary statement

BackgroundGenetic alteration of candidate response elements at their native chromosomal loci is the only valid determinant of their potential transcriptional regulatory activities. Targeted DNA cleavage by Cas9 coupled with cellular repair processes can produce arrays of alleles that can be defined by massively parallel sequencing by synthesis (SBS), presenting an opportunity to generate and survey edited cell populations that include informative alterations. Such editing efforts commonly rely on subclonal enrichment to isolate cells with preferred genotypic properties at target loci; short nucleotide adducts (indices/barcodes) allow PCR-amplified molecules from diverse sample sources to be pooled, sequenced, and demultiplexed to resolve source-specific content. Not widely available, however, are capabilities for barcoding thousands of clones, or for automated analysis of individual candidate regulatory loci PCR-amplified and sequenced from a genetically heterogeneous population--specifically, imputation of discrete genotype(s) by allele definition and abundance, and identification of altered regulatory factor binding motifs.\n\nResultsWe describe a panel of 192 8-nucleotide barcode primers compatible with Illumina(R) sequencing platforms, and the application of these barcodes to genotypic analysis of Cas9-edited clones. Permutations of the ninety-six i7 (read 1) and ninety-six i5 (read 2) barcodes allow unique labeling of up to 9,216 distinct samples. We created three independent Python scripts: SampleSheet.py automates construction of Illumina(R) Sample Sheets encoding up to 9,216 barcode:sample relationships; ImputedGenotypes.py defines alleles and imputes genotypes from demultiplexed fastq files; CollatedMotifs.py flags transcription factor recognition motif matches altered in alleles relative to a reference sequence.\n\nConclusionsCode-enabled definition of alleles and regulatory motifs in sequenced, demultiplexed amplicons facilitates evaluation of genetic diversity in up to 9,216 distinct samples. Here, we demonstrate the utility of three scripts in analysis of cell populations targeted by Cas9 for disruption of glucocorticoid receptor (GR) binding sites near FKBP5, a GR-regulated gene in the human adenocarcinoma cell line A549. SampleSheet.py, ImputedGenotypes.py, and CollatedMotifs.py operate independently and are broadly applicable beyond the case described here.

Standard methods for transgenesis in zebrafish depend on random transgene integration into the genome followed by resource-intensive screening and validation. Targeted vector integration into validated genomic loci using phiC31 integrase-based attP/attB recombination has transformed mouse and Drosophila transgenesis. However, while the phiC31 system functions in zebrafish, validated loci carrying attP-based landing or safe harbor sites suitable for universal transgenesis applications in zebrafish have not been established. Here, using CRISPR-Cas9, we converted two well-validated single insertion Tol2-based zebrafish transgenes with long-standing genetic stability into two attP landing sites, called phiC31 Integrase Genomic Loci Engineered for Transgenesis (pIGLET). Generating fluorescent reporters, loxP-based Switch lines, CreERT2 drivers, and gene-regulatory variant reporters in the pIGLET14a and pIGLET24b landing site alleles, we document their suitability for transgenesis applications across cell types and developmental stages. For both landing sites, we routinely achieve 25-50% germline transmission of targeted transgene integrations, drastically reducing the number of required animals and necessary resources to generate individual transgenic lines. We document that phiC31 integrase-based transgenesis into pIGLET14a and pIGLET24b reproducibly results in representative reporter expression patterns in injected F0 zebrafish embryos suitable for enhancer discovery and qualitative and quantitative comparison of gene-regulatory element variants. Taken together, our new phiC31 integrase-based transgene landing sites establish reproducible, targeted zebrafish transgenesis for numerous applications while greatly reducing the workload of generating new transgenic zebrafish lines. SHORT ABSTRACTTargeted transgenesis into pre-established, "safe harbor," landing sites remains a missing technique in zebrafish. Here, we established phiC31 Integrase Genomic Loci Engineered for Transgenesis (pIGLET) by CRISPR-Cas9-based conversion of two previously validated Tol2 transgenes into attP sites. phiC31-mediated transgenesis into our pIGLET14a and pIGLET24b attP landing sites results in 25-50% germline transmission efficiency and quantifiable transgene activity. Our landing sites are suitable for reproducible, routine zebrafish transgenesis for diverse applications.

RNA-sequencing provides high-dimensional, quantitative measurements of the states of cells, tissues, organs, and whole organisms. Plate-based RNA-seq protocols allow for a wider range of experimental designs than droplet sequencing methods, but are less scalable due to the practical challenges of plate-based liquid handling. Here, we present STOMP-seq, a method that extends SMART RNA-seq protocols, like Smart-seq2 and Smart-seq3, to include sample-identifying barcodes on the 5 end of each amplified transcript. These barcodes allow samples to be pooled immediately after reverse transcription, enabling a 12-fold multiplexing strategy that reduces liquid handling complexity and enzyme costs several-fold. Suitable for both manual and robotic library preparation approaches, STOMP-seq reduces protocol execution times four-fold while improving library complexity and coverage. Together, these advantages combine to make possible new large-scale experimental designs, in particular population-scale sequencing projects like the multi-generational study of gene-expression heritability presented here. STOMP-seq offers a "drop-in" replacement for Smart-seq2 and Smart-seq3, removing practical barriers that currently limit the quality and scope of plate-based transcriptomic data. Graphical Abstract O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=105 SRC="FIGDIR/small/645277v2_ufig1.gif" ALT="Figure 1"> View larger version (52K): org.highwire.dtl.DTLVardef@83f1b4org.highwire.dtl.DTLVardef@717012org.highwire.dtl.DTLVardef@174e34forg.highwire.dtl.DTLVardef@f790b3_HPS_FORMAT_FIGEXP M_FIG C_FIG

The compound eye of Drosophila melanogaster has long been a model for studying genetics, development, neurodegeneration, and heterochromatin. Imaging and morphometry of adult Drosophila and other insects is hampered by the low throughput, narrow focal plane, and small image sensors typical of stereomicroscope cameras. When data collection is distributed among many individuals or extended time periods, these limitations are compounded by inter-operator variability in lighting, sample positioning, focus, and post-acquisition processing. To address these limitations we developed a method for multiplexed quantitative analysis of adult Drosophila melanogaster phenotypes. Efficient data collection and analysis of up to 60 adult flies in a single image with standardized conditions eliminates inter-operator variability and enables precise quantitative comparison of morphology. Semi-automated data analysis using ImageJ and R reduces image manipulations, facilitates reproducibility, and supports emerging automated segmentation methods, as well as a wide range of graphical and statistical tools. These methods also serve as a low-cost hands-on introduction to imaging, data visualization, and statistical analysis for students and trainees.