Molecular Biology

Identifying substrates of protein phosphatases has been technically challenging and has hampered progress in the field of plant sciences. Small molecule inhibitors of protein phosphatases have aided in uncovering classes of phosphatases that target substrates, but that too has severe limitations. Here, we describe a method that enriches phosphorylated substrates using TiO₂ and phospho-tyrosine antibodies in phosphatase knockout lines of Arabidopsis thaliana. When compared to wild-type plants, this approach permits identification of putative substrates and specific phosphorylation sites by mass spectrometry, allowing for further in vitro or functional validation. The key to the approach described here is the use of phosphatase knockout lines to maintain substrates in a phosphorylated state and using phospho-tyrosine antibodies to enrich for tyrosine phosphorylated peptides.

Gastric cancer remains a major global health challenge, and reliable prognostic biomarkers are urgently needed to guide treatment decisions. Here, we present a simple and efficient protocol for a novel liquid biopsy approach based on quantifying gastric fluid DNA (gfDNA) collected during routine esophagogastroduodenoscopy (EGD). We have previously shown that gfDNA carries gastric cancer–derived mutations; moreover, its concentration increases with tumor progression and varies according to cancer prognosis. This empirically observed increase in gfDNA may mechanistically stem from enhanced cellular turnover, tissue disorganization, dysbiosis of the local microbiota, and/or fluctuations in immune cell infiltrates. Surprisingly, however, in patients diagnosed with gastric cancer, elevated gfDNA levels were also associated with improved survival. This paradoxical finding may be reconciled by an increased anti-tumor immune cell response in treatment-responsive gastric cancers, as well as by the contribution of non-tumoral DNA from inflammatory processes within the microenvironment of the stomach. Here, we detail a standardized protocol for gastric fluid collection and processing, designed to support downstream gfDNA quantification among other potential molecular applications.

Chromatin modifications regulate genome function by recruiting proteins that control transcription, genome organization, and DNA repair. Identifying the proteins associated with specific chromatin modifications is therefore essential for understanding how these regulatory processes operate. Traditional approaches, including chromatin immunoprecipitation and affinity purification coupled to mass spectrometry, have uncovered many chromatin-associated proteins. However, they often rely on crosslinking and chromatin fragmentation, which can disrupt native chromatin architecture and limit the detection of transient interactions. Here, we describe a proximity-labeling protocol for identifying the chromatin-dependent protein interactome associated with specific chromatin marks, termed ChromID. ChromID uses engineered chromatin readers (eCRs) fused to a promiscuous biotin ligase, which labels proteins in the immediate vicinity of the targeted chromatin mark. The protocol includes in vivo biotin labeling, nuclear extract preparation, streptavidin-based enrichment, and tryptic digestion for downstream LC-MS/MS analysis. The protocol has been validated across multiple cell types and chromatin contexts and can be extended to other chromatin-associated proteins, providing a versatile approach to profile chromatin-associated proteomes within their native cellular environment.

RNA modifications and their “writer,” “eraser,” and “reader” proteins are emerging as key regulators of gene expression and DNA repair through dynamically regulating RNA:DNA hybrids, or R-loops, during transcription. Therefore, it is paramount to develop rigorous techniques for accurate analysis of R-loop modifications. A convenient method for analyzing RNA modifications within total RNA is by dot blot with specific RNA modification antibodies; however, analysis of the modification of the RNA moiety within R-loops presents specific challenges. Here, we provide a detailed protocol for the production or purification of DNA containing R-loops in vitro and from cells, and the analysis of the RNA moiety modifications by dot blot. The DNA containing R-loops is treated with either mock or RNase H, which specifically degrades the RNA within RNA:DNA hybrids, to control for the specificity of the signal as originating from R-loops. Known quantities of the mock or RNase H–treated DNA are then spotted on three membranes, each blotted with antibodies that recognize double-stranded DNA, RNA:DNA hybrids, or the specific RNA modification antibodies of interest, such as m⁶A or ac⁴C. Thus, this protocol is useful to both biochemists and cell biologists with scientific interests at the intersection of R-loops and epitranscriptomics.

Riboswitches are structured non-coding RNA elements that regulate gene expression in response to small molecules; they serve as valuable systems in both public health and biophysical research by elucidating principles around RNA–ligand interactions, structure, and cellular function. Traditional approaches to studying riboswitches have relied on low-throughput techniques such as reporter assays or gel electrophoresis analysis of transcriptional products, which are limited in scalability. In this study, we present a high-throughput protocol to characterize the transcriptional activity of nearly 2,000 natural variants of the fluoride riboswitch in in vitro transcription. Starting with bioinformatics, we compiled a comprehensive dataset of riboswitch variants and then employed massive parallel oligonucleotide synthesis to generate an oligo pool of the riboswitch library. This pool was transcribed in vitro, converted into an Illumina-compatible next-generation sequencing (NGS) library, and analyzed to identify transcriptionally active riboswitch candidates. The workflow integrates natural riboswitch bioinformatic acquisition into a quantitative readout in a single streamlined pipeline, enabling large-scale exploration of transcriptional riboswitch function. This protocol offers a scalable method for mapping genotype-to-function relationships across transcriptional riboswitch families, accelerating the identification of functional variants for desired applications.

Genetically encodable reporters that produce signals detectable in deep tissues offer a powerful tool for noninvasive monitoring of molecular events in vivo. Although magnetic resonance imaging (MRI) is a standard technique for noninvasive clinical imaging, its wider application in detecting molecular activities has been constrained by the lack of programmable sensors. This limitation is in stark contrast to the widespread use of fluorescent reporter–derived sensors in cultured cells and in transparent specimens. To overcome this limitation, we recently developed the modular aquaporin-based protease-activatable probe for enhanced reporting (MAPPER) platform. This sensor engineering framework integrates a metal-free MRI reporter derived from human aquaporin-1 (hAqp1) with synthetic protease-based circuits. This integration facilitates the modular and scalable creation of a wide range of sensors by regulating protease activity through precise molecular events, such as protein–protein interactions, pharmacological inhibition, and second messenger signaling. In this paper, we present a detailed protocol for constructing and deploying sensors using the MAPPER paradigm. The protocol encompasses genetic design, lentiviral production, stable cell line generation, biochemical and microscopic validation of sensor function, diffusion-weighted MRI, and MR image analysis to quantify sensor signals in terms of the apparent diffusion coefficient. We describe two distinct MAPPER architectures: DD-MAPPER, which leverages protease-controlled protein degradation, and ER-MAPPER, which utilizes protease-controlled, subcellular trafficking. The MAPPER framework allows adaptation to various molecular targets without the need to redesign the core MRI reporter mechanism, making MAPPER a versatile platform for noninvasive biosensing in living cells and tissues.

Chemotherapy-resistant persister cells are a major driver of cancer recurrence, yet their epigenetic basis remains poorly characterized. This protocol describes a computational pipeline for identifying DNA-binding factors (DBFs) that are enriched in accessible chromatin that collectively define a persister cell signature (PCS). Starting from single-nucleus ATAC-seq (snATAC-seq) data processed through the 10x Genomics CellRanger ARC pipeline, this protocol covers (1) the creation of a Seurat/Signac object with ATAC peaks, (2) the optional integration of DNA-binding data from the ReMap2022 database as a per-cell chromatin module assay, (3) differential accessibility analysis across clinically defined comparison groups, and (4) identifying and defining the top enriched DBFs as the PCS. This approach is applicable to any snATAC-seq dataset in which cells can be grouped by clinical response, treatment status, or resistance phenotype.

Site-directed mutagenesis is an indispensable molecular biology tool, but traditional methods often suffer from extended reaction time, structural limitations, and variable success rates. This article details three optimized protocols: P3a (primer pairs with 3′-overhangs, version a), P3b, and QuickChange 2.0, which rely on two highly processive DNA polymerases (Platinum SuperFi II and Q5) to accelerate and standardize plasmid engineering. The P3a method utilizes partially complementary primer pairs with distinct 3′-overhangs, achieving ~100% efficiency and enabling seamless cassette mutagenesis (insertion, deletion, and replacement). Building on this, the P3b method introduces specific thermal cycling modifications and a pre-denaturation step to overcome structural barriers resulting from GC-rich sequences. QuickChange 2.0 applies these two advanced polymerases to completely complementary primer pairs, even though the average efficiency decreases to 50%–60%. Replacing Pfu with the highly processive DNA polymerases also reduces PCR time to approximately 2 h. Thus, these new methods are more efficient and rapid than classical QuickChange mutagenesis based on Pfu polymerase.

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 cap-independent 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 translation activity under controlled conditions and can additionally be applied to evaluate mRNA modifications designed to enhance translation efficiency.

Mosaic animals are highly valuable for investigating complex biological processes and cell lineages in vivo. Traditional mosaic techniques in Drosophila, such as the FRT/Flp system, rely on exogenous site-specific recombination sequences, preventing their application to unmodified mutant chromosomes or wild-derived strains. Mosaic analysis by gRNA-induced crossing-over (MAGIC) overcomes this limitation by utilizing the CRISPR/Cas9 system to generate targeted double-strand breaks (DSBs) that induce somatic homologous recombination in precursor cells. Here, we describe a comprehensive protocol for applying MAGIC with a newly developed, genome-wide MAGIC kit. This protocol utilizes optimized gRNA-markers with the Qtg2.1 scaffold for high-efficiency clone induction, alongside improved fluorescent labeling strategies for both positive MAGIC (pMAGIC) and negative MAGIC (nMAGIC). The procedure details the genetic crossing schemes, temporal induction of clones, and tissue processing for diverse Drosophila cell types. This method enables convenient mosaic analysis across all chromosomes and allows for the study of pericentromeric genes, deficiency chromosomes, and species-specific alleles in interspecific hybrids.