Current Protocols

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.

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.

DNA secondary structures are essential elements of the genomic landscape, playing a critical role in regulating various cellular processes. These structures refer to G-quadruplexes, cruciforms, Z-DNA or H-DNA structures, amongst others (collectively called non-B DN), which DNA molecules can adopt beyond the B conformation. DNA secondary structures have significant biological roles, and their landscape is dynamic and can rearrange due to various factors, including changes in cellular conditions, temperature, and DNA-binding proteins. Understanding this dynamic nature is crucial for unraveling their functions in cellular processes. Detecting DNA secondary structures remains a challenge. Conventional methods, such as gel electrophoresis and chemical probing, have limitations in terms of sensitivity and specificity. Emerging techniques, including next-generation sequencing and single-molecule approaches, offer promise but face challenges since these techniques are mostly limited to only one type of secondary structure. Here we describe an updated version of a technique permanganate/S1 nuclease footprinting, which uses potassium permanganate to trap single-stranded DNA regions as found in non-B structures, in combination with S1 nuclease digest and adapter ligation to detect genome-wide non-B formation. To overcome technical hurdles, we combined this method with direct adapter ligation and sequencing (PDAL-Seq). Furthermore, we established a user-friendly pipeline available on Galaxy to standardize PDAL-Seq data analysis. This optimized method allows the analysis of many types of DNA secondary structures that form in a living cell and will advance our knowledge of their roles in health and disease.

Evaluating cellular responses to toxic compounds is essential for assessing the safety and potential hazards of drugs, environmental pollutants, and food contaminants. Traditional in vitro methods often lack the precision and scalability required for comprehensive toxicological assessments. This protocol presents an efficient and scalable approach for quantifying cellular damage and toxicity using organ-on-chip technology, specifically the OrganoPlate(R) platform. By combining fluorescent probes--DRAQ7 for cell membrane integrity, ActinGreen for cytoskeletal changes, and NucBlue for nuclear counting--with high-throughput image analysis via CellProfiler, this method provides detailed and quantitative insights into cellular damage induced by toxic compounds. The protocol includes step-by-step instructions for staining, image acquisition, and data analysis, as well as troubleshooting guidance. CellProfilers open-source nature, flexibility, and automation capabilities enable reproducible, high-throughput workflows, offering significant advantages over traditional manual image analysis. Its ability to assess cytotoxicity in human tissue models makes this protocol a valuable tool for safety testing in drug discovery and environmental toxicology. Furthermore, the approach is highly adaptable, accommodating a variety of cell types and toxic compounds, and is well-suited for rapid screening and risk assessment across diverse research and industrial applications.

Eukaryotic proteins can be expressed in different heterologous systems. However, mammalian proteins in general have specific post-translational processing requirements that may not be fulfilled by a regular bacterial expression system. In this study, we use an insect cell system to express a mammalian protein of interest. Spodoptera frugiperda (Sf21) cells were used in conjunction with a baculoviral expression system to produce the cytosolic domain (CD) of IRE1, an endoplasmic reticulum (ER) stress sensor protein. Inositol Requiring Enzyme 1 (IRE1) is a dual function kinase and endoribonuclease protein that cleaves X-box binding protein (XBP1) mRNA. We used the pFastBac plasmid to insert the coding sequence into a recombinant bacmid shuttle vector which was then used to infect Sf21 cells. The expressed protein was then purified with an MBPTrap column to obtain >85% pure protein.

Intricate interactions between RNA-binding proteins (RBPs) and RNA play pivotal roles in cellular homeostasis, impacting a spectrum of biological processes vital for survival. UV crosslinking methods to study protein-RNA interactions have been instrumental in elucidating their interactions but can be limited by degradation of target proteins during the process, low signal-to-noise ratios, and non-specific interactions. Addressing these limitations, we describe reCRAC (reverse CRAC), a novel adaptation of the CRAC (crosslinking and analysis of cDNA) technique, optimized for yeast Saccharomyces cerevisiae. Like CRAC, reCRAC applies tandem affinity purification to yield highly enriched protein preparations. However, reCRAC is redesigned to work with unstable proteins. This is achieved by lysing the cells directly into highly denaturing buffer conditions, followed by stringent purification steps. The reCRAC method was successfully applied to the easily degraded yeast protein Pin4, allowing identification of precise binding sites at base-pair resolution with greatly reduced target protein degradation and improved signal-to-noise ratios.

pomBseen is a image analysis pipeline for the quantitation of fission yeast micrographs containing a brightfield channel and up to two fluorescent channels. It accepts a wide range of image formats and produces a table with the number, size and total and nuclear fluorescent intensities of the cells in the image. Written in MATLAB, pomBseen is also available as a standalone application.

Transfer RNA (tRNA), its post-transcriptional modifications, and tRNA-derived fragments (tRFs) play essential roles in cellular processes and gene regulation. Here, we present a fast and efficient tRNA isolation using silica spin columns. To analyze the isolated tRNA and detect tRFs, we describe a sensitive and cost-effective non-radioactive Northern blotting technique. Additionally, this blotting method is compatible with chemical affinity modifiers, such as [p-(N-acrylamino)-phenyl]mercuric chloride (APM) or 3-(acrylamido)phenylboronic acid (APB) enabling the detection of chemical modifications in specific tRNA isoacceptors. Graphical abstract O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=146 SRC="FIGDIR/small/681074v1_ufig1.gif" ALT="Figure 1"> View larger version (39K): org.highwire.dtl.DTLVardef@1b0618forg.highwire.dtl.DTLVardef@d507a4org.highwire.dtl.DTLVardef@1e8b6faorg.highwire.dtl.DTLVardef@1450326_HPS_FORMAT_FIGEXP M_FIG C_FIG HighlightsO_LIRapid (<30 min) tRNA isolation from total RNA using silica containing spin columns C_LIO_LIProcedure for DNA probes biotinylation via terminal deoxynucleotidyl transferase C_LIO_LINon-radioactive chemiluminescent Northern blotting C_LI

Image analysis is a rapidly developing field that provides unique opportunities to characterize and quantify spatial information in images. We study cis-regulatory modules (CRMs), non-coding DNA regions that regulate gene expression, during development using fluorescent reporters in vivo. Characterizing CRM activity during development presents challenges including image segmentation into biologically relevant regions of interest that are not easily distinguishable via common segmentation methods, fluorophore band passing, and variable transfection undermine standardized analysis. To quantify and compare CRM activity levels, we compiled an open-source computer vision tool stack in the form of a Python-based Jupyter notebook and tested four analysis methods to assess their efficacy in quantifying CRM expression in limb development. This Jupyter notebook provides a reproducible, standardized workflow that can be adapted to numerous image analysis applications.

Polarized epithelial cells form a tightly packed monolayer where individual cells are connected by cell-cell junctions, including tight junctions (TJ) and adherens junctions (AJ). Here, we present techniques for quantifying the horizontal and vertical distribution of junctional proteins in confluent, fixed epithelial cells. This approach is utilized to evaluate variations in the intensity and localization of the proteins that compose the AJ and TJ under different experimental conditions. Although our protocol is optimized for Madin-Darby Canine Kidney (MDCK) cells, it is adaptable to any cell line capable of forming cell-cell junctions. For complete details on the use and execution of this protocol, please refer to Rabino et al. (2024). 1 Graphical abstract O_FIG O_LINKSMALLFIG WIDTH=190 HEIGHT=200 SRC="FIGDIR/small/651687v1_ufig1.gif" ALT="Figure 1"> View larger version (53K): org.highwire.dtl.DTLVardef@1d1db4forg.highwire.dtl.DTLVardef@954ea1org.highwire.dtl.DTLVardef@5d43eborg.highwire.dtl.DTLVardef@1162077_HPS_FORMAT_FIGEXP M_FIG C_FIG

Genetic modifications using CRISPR-Cas9 have revolutionized cancer research and other pre-clinical studies. Exceptionally, these efficient tools are inadequate in a few disease models and cell lines due to the aberrant differentiation states and the accumulation of excessive somatic mutations that compromise the robustness of viral gene delivery and stable transduction. A couple of B lymphoma cell lines fall into this category where lentiviral transfection becomes inefficient and exhibits variable efficiency. Additionally, lentiviral delivery requires high biosafety levels. To address this challenge, we have developed a two-step strategy that supports CRISPR-Cas9 through lentivirus and murine ecotropic {gamma}-retrovirus. By engineering B lymphoma cell lines to express Cas9 and mCat-1, a specific receptor for ecotropic retroviruses, we enable efficient and safe gene editing through ecotropic {gamma}-retrovirus. We demonstrate the efficacy of this method by generating IgM-deficient B lymphoma cell lines. This innovative approach simplifies protocols, enhances accessibility, and paves the way for standardized gene manipulation of B cell lymphoma models for molecular cell biology research.

Investigating the RNA regulation landscape primarily relies on our understanding of how RNA-protein interactions are governed in various cell types, including neurons. Analysis of RNA-protein interactions in physiological environments warrants the development of new tools that rely on RNA manipulation. Recently, A CRISPR-based RNA-editing tool (dCas13b-ADAR2DD) was developed to mitigate disease associated point mutations in cell lines. Here, we have explored the targeted sequence editing potential of the tool (dCas13b-ADAR2DD system) by adapting it to manipulate RNA function with an aim to visualize RNA editing in primary hippocampal neurons. This is a two-component system that includes a programmable guide RNA (gRNA) complementary to the target RNA, and a catalytically dead version of the Cas13b enzyme fused to ADAR. The RNA editing protocol outlined in this manuscript relies on using the gRNA-dependent targeting of dCas13b-ADAR fusion protein to the mutant form of mDendra transcript. We first abrogated the fluorescence of Dendra2 by introducing a nonsense mutation that precludes the formation of the functional protein. To visualize the efficacy of the RNA editing in neurons, we used the dCas13b-ADAR2DD system to edit specific nucleotides within the Dendra2 mRNA to restore the amino acid codes critical for Dendra2 fluorescence. This method therefore lays the foundation to future studies on the dynamicity of activity-induced RNA-protein interactions in neurons and can be extended to manipulate the endogenous RNome in diverse neuronal subtypes. Furthermore, this methodology will enable investigators to visualize the spatial and temporal resolution of RNA-protein interactions without altering the genomes via conventional methods. HighlightsO_LICRISPR-Cas13 based application that enables site specific A-to-I in target RNAs directed by gRNA; optimized in neurons. C_LIO_LIEnables the temporal mapping of developmentally relevant RNAs and their cis-interacting RNA binding proteins. C_LI

Optogenetic gene therapy may employ recombinant adeno-associated virus (AAV) vectors for specific preclinical applications in the treatment of inherited retinal diseases. Human retinal explants as an ex vivo model assist determine cellular tropism and the level of subsequent gene expression followed by delivery of viral vectors. Extracts of retinal tissue can be obtained and cultured for a certain time; thus, it is possible to transduce and confirm the effectiveness of the virus using various methods, including immunohistochemical staining. Consequently, we described method for harvesting and culturing human retinal fragments from divergent parts of the retina that differ anatomically and molecularly. The fragments were fixed, sectioned and stained. A dissociation method has also been described to obtain a retinal cell suspension that can be used in other experiments such as flow cytometry or mRNA/protein level expression assays.

The ultimate output of gene expression is to ensure that proteins are synthesized at the right levels, locations, and timings. Recently different imaging-based methods have been developed to visualize the translation of single mRNA molecules. These methods rely on signal amplification with the introduction of an array of a short peptide sequence (a tag such as SunTag), recognized by a genetically encodable single-chain antibody (a detector such as scFv). In this chapter, we discuss such methods to image and quantify translation dynamics in the early Drosophila embryo and provide examples based on a twist-32XSunTag reporter. We outline a step-by-step protocol to light-up translation in living embryos. We also detail a combinatorial strategy in fixed samples (smFISH-IF), allowing to distinguish single mRNA molecules engaged in translation.

i.Accurate identification of the locations of endogenous proteins is crucial for understanding their functions in tissues and cells. However, achieving precise cell-type-specific labeling of proteins has been challenging in vivo. A notable solution to this challenge is the self-complementing split green fluorescent protein (GFP1-10/11) system. In this paper, we present a detailed protocol for labeling endogenous proteins in a cell-type-specific manner using the GFP1-10/11 system in fruit flies. This approach depends on the reconstitution of the GFP1-10 and GFP11 fragments, creating a fluorescence signal. We insert the GFP11 fragment into a specific genomic locus while expressing its counterpart, GFP1-10, through an available Gal4 driver line. The unique aspect of this system is that neither GFP1-10 nor GFP11 alone emits fluorescence, enabling the precise detection of protein localization only in Gal4-positive cells expressing the GFP11 tagged endogenous protein. We illustrate this technique using the adhesion molecule gene teneurin-m (Ten-m) as a model, highlighting the generation and validation of GFP11 protein trap lines via Minos-mediated integration cassette (MiMIC) insertion. Furthermore, we demonstrate the cell-type-specific labeling of Ten-m proteins in the larval brains of fruit flies. This method significantly enhances our ability to image endogenous protein localization patterns in a cell-type-specific manner and is adaptable to various model organisms beyond fruit flies.

An often-overlooked aspect of mitochondrial biology is the mitochondrial DNA (mtDNA). The multi-copy mtDNA is highly dynamic, with changes in supercoiling, synthesis rates, and turnover rates that are tightly associated with mitochondrial and cellular functions. To better understand the state of the mtDNA, here we describe a protocol that selectively incorporates bromodeoxyuridine into the mtDNA for subsequent measurement via an adapted Southern blot followed by immunoblotting (a.k.a. Southwestern blot). This basic protocol can be applied with slight modifications for the measurement of mtDNA synthesis, turnover or supercoiling to understand mtDNA changes.

Studying adipogenesis and adipocyte biology requires the isolation of primary preadipocytes from adipose tissues. However, primary preadipocytes have a limited lifespan, can only undergo a finite number of divisions, and often lose their original biological characteristics before becoming senescent. The repeated isolation of fresh preadipocytes, particularly from young pups or aged animals, is costly and time-consuming. Immortalization of these cells offers a solution by overcoming cellular senescence and maintaining proliferative capacity, allowing for long-term studies without the continuous need to isolate new cells from animals. Immortalized cell lines thus provide a consistent and reproducible experimental model, significantly reducing variability across different animals. However, successfully establishing immortalized preadipocyte cell lines presents challenges, including selecting appropriate adipose tissue depots, isolating primary preadipocytes, and choosing an effective immortalization strategy. In this study, we present optimized protocols and share first-hand experiences establishing immortalized brown and white preadipocyte cell lines from young and aging mice. These protocols offer a valuable resource for researchers studying adipogenesis, metabolism, and adipocyte biology. Support Protocol 1: Retrovirus production Basic Protocol 1: Isolation and culture of primary brown and white preadipocytes from mouse interscapular brown adipose tissue (iBAT) and subcutaneous white adipose tissue (sWAT) in the same region Basic Protocol 2: Immortalization of mouse brown and white preadipocytes Basic Protocol 3: Selection of immortalized preadipocytes Basic Protocol 4: Selection of single-cell clones of immortalized preadipocytes Support Protocol 2: Cryopreservation of immortalized preadipocytes Support Protocol 3: Wake up and culture of immortalized preadipocytes Support Protocol 4: Subculture and expansion of immortalized preadipocytes Basic Protocol 5: Differentiation of immortalized mouse brown and white preadipocytes Support Protocol 5: Lipid droplet staining and nucleus counterstaining Support Protocol 6: Mitochondria staining and nucleus counterstaining

In less than a year, SARS-CoV-2 (SARS2) has managed to displace Mycobacterium tuberculosis (Mtb) as the leading cause of death worldwide due to a single infectious agent. Both pathogens affect the respiratory tract, mainly the lungs. However, the impact that a possible Mtb + SARS2 co-infection can have on the host response is still unknown. Herein we propose (depict) a rigorous system to evaluate the complex interaction between the two infections in vitro in a lung epithelial cell line (A549). Overall, the process includes eight steps: (I) Mtb culture, (II) cell maintenance, (III) preparation of viral stocks, (IV) determination of infectious titers, (V) Mtb and SARS2 co-infection, (VI) determination of intracellular bacterial load, (VII) SARS2 viability test, and (VIII) decontamination of supernatants. This comprehensive protocol will allow experimentalists to study the pathogenesis of co-infection in vitro, and facilitate collaborative work in the literature.

We describe a method for the generation and deep imaging of human precision-cut lung slices (PCLS). PCLS bridge the gap between in vivo and in vitro studies, providing a robust system for visualizing events from tissue to subcellular levels in the three-dimensional lung environment, with the preservation of all resident cell types and cell-cell interactions. They also constitute a validated model for studying host cell-pathogen interactions. Here, we detail the generation of human PCLS, followed by their infection and imaging by laser scanning confocal microscopy and transmission electron microscopy (TEM). We establish the conditions for ex vivo infection and replication of two pathogens of relevance to human respiratory health: a virus (SARS-CoV-2) and a bacterium (Mycobacterium tuberculosis, Mtb). PCLS can be obtained in a single day, infected the next day, and were successfully cultivated for up to a week in this study. Imaging was performed on fixed samples. The preparation of PCLS took one day for confocal imaging and five days for TEM imaging. All procedures are readily adaptable to explore other pathogens and other species and are easy to implement by users with experience in tissue culture. Some specialist equipment (an Alabama tissue slicer) is required for PCLS generation.

Whole-mount in situ hybridization (WISH) is a powerful and widely used technique to visualize the expression pattern of genes in different biological systems. Here we describe a new protocol for ISH and immunostaining in the planarian Schmidtea mediterranea. The new Nitric Acid/Formic Acid (NAFA) protocol is compatible with both assays and prevents degradation of the epidermis or blastema. Instead of proteinase K digestion, formic acid treatment is used to permeabilize tissues and preserve antigen epitopes. We show that the NAFA protocol successfully permits development of chromogenic and fluorescent signals in situ, while preserving the anatomy of the animal. Further, the immunostaining of different proteins was compatible with the NAFA protocol following fluorescent in situ hybridization. Finally, we demonstrate with high resolution confocal imaging that the regeneration blastema is preserved when using the new method. This new NAFA protocol will be a valuable technique to study the process of wounding response and regeneration.