Molecular Biology

Human tissue samples represent the gold standard for obtaining clinically relevant and translatable insight into disease processes that in vitro systems cannot fully reproduce. However, patient-derived samples are often limited in size and availability, limiting the number of downstream assays that can be performed. To maximize the use of invaluable human samples, we present a protocol for the tandem extraction of high-quality RNA and protein from the same tissue section. This method has been optimized for 15–30 mg tissue sections, enabling more experimental conditions and technical replicates, while minimizing intrasample variability associated with heterogeneous tissues. This protocol also avoids potentially hazardous solvents present in phenol-chloroform-based methods such as TRIzol, providing a safer and more accessible workflow without compromising biomolecule integrity. This protocol was developed and validated using atherosclerotic plaque tissue from carotid endarterectomy, a very challenging tissue type to work with due to extensive calcification, necrosis, and limited surgical availability. We have also validated this method using mouse aortic tissue and cultured THP-1 cells, demonstrating its versatility across sample input types. As this protocol relies on standard column-based RNA extraction kits and commonly available reagents for protein precipitation and extraction, this methodology is widely accessible and easy to implement as a standard, streamlined workflow.

Transcription factors (TFs) regulate gene expression by binding to cis-regulatory elements in the genome. Understanding transcriptional regulation requires genome-wide characterization of TF occupancy across different chromatin contexts, yet simultaneous assessment of TF binding for multiple factors remains technically challenging. Here, we describe a detailed and reproducible protocol for cFOOT-seq, a cytosine deaminase–based genomic footprinting assay by sequencing, which enables antibody-independent, base-resolution profiling of chromatin accessibility, nucleosome organization, and TF occupancy. In cFOOT-seq, the double-stranded DNA (dsDNA) cytosine deaminase SsdA_tox converts cytosine to uracil in accessible chromatin, whereas TF binding and nucleosome occupancy locally protect DNA from deamination. Using the FootTrack analysis framework, deamination patterns generated by cFOOT-seq are quantitatively analyzed to derive standardized footprint and chromatin organization profiles at base resolution across the genome. Because cFOOT-seq preserves genomic DNA integrity during deamination-based footprinting, it is compatible with ATAC-seq-based chromatin enrichment. ATAC-combined implementations reduce sequencing depth requirements and improve scalability for footprint-focused analyses, supporting applications in low-input and single-cell settings. This protocol provides a practical framework for genome-wide TF footprint profiling and can be readily applied to dissect gene regulatory mechanisms in development, immunity, and disease, including cancer.

Extrachromosomal circular DNA (eccDNA) is a type of circular DNA that exists independently of chromosomes and has garnered significant attention in various fields, particularly in the context of smaller eccDNAs, which have considerable roles in gene regulation through various mechanisms. Current methods such as Circle-Seq and 3SEP can enrich small eccDNAs during sample preparation, but most bioinformatics pipelines remain challenging, exhibiting low accuracy and efficiency. This protocol describes the detailed workflow of a newly developed bioinformatics analysis pipeline, named EccDNA Caller based on Consecutive Full Pass (ECCFP), to accurately identify eccDNA from long-read Nanopore sequencing data. Compared to other pipelines, ECCFP significantly improves detection sensitivity, accuracy, and runtime efficiency. The process includes raw data quality control, trimming of adapters and barcodes, alignment to a reference genome, and identification of eccDNA, with detailed results encompassing accurate positioning of eccDNA, consensus sequences, and variants of individual eccDNA.

Agrobacterium rhizogenes–mediated hairy root transformation provides a rapid platform for gene function analysis prior to stable whole-plant transformation. However, most existing hairy root transformation methods rely on tissue culture and require chemical or fluorescence-based selection, which increases experimental complexity. Here, we describe a tissue culture–free soybean hairy root transformation protocol incorporating the RUBY visual reporter system. While this work does not introduce a new transformation concept, it presents a streamlined implementation of established soybean hairy root methodologies that emphasizes procedural simplicity, reduced handling, and faster access to functional root material. Transgenic roots expressing RUBY can be directly identified by red pigmentation with the naked eye. In RUBY-positive roots, candidate genes driven by the CaMV 35S promoter showed higher expression levels than those in empty-vector controls, indicating that the system supports effective gene expression. Using this procedure, clearly identifiable transgenic hairy roots can be obtained within 20 days. Overall, this protocol simplifies induction and screening while reducing operational complexity and equipment requirements.

In wheat and other crops, some genes display presence/absence variation, and it is occasionally beneficial to select for the absent allele to remove a functional gene. However, current high-throughput genotyping methods used to detect the absence of genes tend to be inconsistent and inconclusive. Kompetitive allele-specific PCR (KASP) and PCR allele competitive extension (PACE) are two well-established methods for allele-specific polymerase chain reaction (AS-PCR) assays, each using fluorescence resonance energy transfer (FRET) to generate a signal for each allele, typically targeting biallelic single-nucleotide polymorphisms. KASP and PACE methods are more difficult to apply to alleles with presence/absence variation because the lack of amplification of the absent allele is indistinguishable from a failed PCR. Here, we present a multiplex fluorescence-based absent allele–specific amplification (FAASA) method using the PACE marker system (compatible with KASP markers) to detect the absence of one particular or all alleles of a target sequence using a primer mix consisting of one target-specific primer pair (TSP) and a second primer set specific to a highly conserved endogenous gene known as a core gene–specific primer pair (CGSP). The forward primer of each pair is tagged with a 5′ terminal tail complementary to dye-labeled oligonucleotides in commercially available FRET cassettes. Lines that amplify only the core gene do not carry the target, while lines that amplify both the core gene and the target carry alleles of both the core gene and the target. The inclusion of the CGSPs allows researchers to confidently distinguish lines with absent alleles of the target from lines with failed PCR reactions, which can happen due to various reasons, including inadequate DNA quality or quantity.

The deletion and mutation of Topoisomerase 3β (TOP3B) is linked to multiple neurological disorders and is the only known topoisomerase that is also catalytically active on RNA in vitro and in cells. Uniquely, TOP3B is primarily localized to the cytoplasm, binds to open reading frames of mRNA, and regulates mRNA stability and translation in a transcript-specific manner. A common approach for studying TOP3B activity in cells is immunodetection of TOP3B•RNA covalent intermediates after bulk RNA isolation. However, in this approach, the RNA species is unknown and is not selective for the major TOP3B substrate, mRNA. In this protocol, we describe a recently developed and optimized protocol for capturing TOP3B•mRNA covalent intermediates using oligo-dT isolation of mRNA under protein-denaturing conditions. Covalent intermediates are then detected by a dual membrane slot blotting strategy with nitrocellulose and positively charged nylon membranes. Nitrocellulose membrane-bound TOP3B•mRNA covalent intermediates are analyzed by immunodetection, and nylon membrane-bound free mRNA is stained with methylene blue. The protocol detailed below has been validated with wildtype and mutant 3xFLAG-tagged TOP3B expressed in Neuro2A cells, with additional optimization for slot blotting using recombinant EGFP.

Amphibian retinas contain “green” rods, which are rod-shaped photoreceptors with a cone-type visual pigment. These rods are considered a potentially transitional photoreceptor type, but their phototransduction cascade’s molecular composition has remained uncertain. Here, we present a streamlined electrophysiology-molecular workflow that enables the rapid spectral identification, physical capture, and targeted single-cell reverse transcription-polymerase chain reaction (RT-PCR) of individual amphibian photoreceptors. After suction-pipette spectral screening under alternating red and green illumination, electrophysiologically identified cells are isolated and processed directly for reverse transcription and PCR. Coupling real-time functional phenotyping with sensitive molecular profiling provides a practical tool for resolving photoreceptor molecular heterogeneity and investigating evolutionary transitions between rod and cone phenotypes.

RNA-binding proteins (RBPs) have pleiotropic roles in modulating the physiology of both eukaryotic and prokaryotic cells, enabling them to adapt to environmental variations. The importance of RBPs has led to the development of a variety of methods aiming to identify them. However, most of these approaches have primarily been implemented and optimized in eukaryotic systems. To both uncover novel RBPs involved in Bacillus subtilis sporulation and capture their RNA-binding ability dynamically, we adapted the orthogonal organic phase separation technique (OOPS), which had previously been used in Escherichia coli to reveal its RNA-binding proteome (RBPome). We optimized the UV cross-linking process used to stabilize RNA–protein interactions in vivo and the bacterial lysis process to overcome the robust cell wall of Gram-positive sporulating cells. RNA–protein complexes are then recovered after phase separation steps using guanidinium thiocyanate–phenol–chloroform, and RNA-associated proteins are identified and label-free-quantified by liquid chromatography–mass spectrometry. Collecting samples at various time points during sporulation further enables tracking the dynamics of the RBPome. In addition to being applicable to bacteria and requiring minimal starting material, this method has provided a comprehensive map of the RBPome during sporulation, refining the roles of known factors and revealing new players.

ADGRL4 is an adhesion G protein–coupled receptor (aGPCR) implicated in multiple tumours. In our experience, conventional insect cell-based baculovirus expression systems have not yielded sufficient correctly folded ADGRL4 protein for purification and cryo-electron microscopy (cryo-EM) analysis. Here, we describe aGPCR-HEK, a six-week protocol that establishes stable tetracycline-inducible mammalian HEK293S GnTI- TetR cell lines expressing N-terminally HA- and GFP-tagged aGPCRs. The method comprises lentiviral production in Lenti-X 293T cells, transduction of target adherent HEK293S GnTI- TetR cells, flow cytometry enrichment of uninduced GFP-positive cells displaying leaky expression, adaptation to suspension culture, and large-scale tetracycline induction and harvesting of cells for downstream purification and cryo-EM. The system yields reproducible, milligram-scale quantities of folded aGPCR suitable for structural and biochemical studies.

RNA-binding protein (RBP)–RNA interactions are fundamental for gene regulation and cellular homeostasis. Ataxin-2 is an RBP that has been shown to play an instrumental role in pathophysiological processes by binding to mRNA. Methods such as RNA immunoprecipitation (RIP), cross-linking immunoprecipitation (CLIP), and their variants can be used to study the interactions between Ataxin-2 and its targets, although their high sample requirements and labor-intensive workflows can limit their widespread use. RNA editing-based approaches, such as targets of RBPs identified by editing (TRIBE), provide effective alternatives. TRIBE enables transcriptome-wide identification of RBP targets by inducing site-specific adenosine-to-inosine (A-to-I) editing, which is subsequently detected through high-throughput RNA sequencing in both in vivo and in vitro systems. Compared to in vivo models, cell lines offer a rapid and flexible experimental design. Drosophila S2 cells are a commonly used insect cell line to investigate RNA–protein dynamics and serve as a versatile platform for studying RBP function. Here, we describe a protocol used for identifying RNA targets of Ataxin-2, a versatile RBP involved in post-transcriptional and translational regulation, in S2 cells using TRIBE. This method allows rapid, efficient, and reliable identification of Ataxin-2-associated RNA targets and can be readily applied to other RBPs.