Molecular Biology

Laboratory-developed tests (LDTs) are optimal molecular diagnostic modalities in circumstances such as public health emergencies, rare disease diagnosis, limited budget, or where existing commercial alternatives are unavailable, limited in supply, or withdrawn, either temporarily or permanently. These tests reduce access barriers and enhance equitable clinical practice and healthcare delivery. Despite recommendations for the development of nucleic acid amplification tests, procedural details are often insufficient, inconsistent, and arbitrary. This protocol elucidates the methodology used in the development of a fully automated real-time polymerase chain reaction (qPCR)-based test, using the Panther Fusion^® Open Access^TM functionality, for the detection of Streptococcus agalactiae in pregnant women, using selectively enriched rectovaginal swabs. In addition, guidelines are provided for oligonucleotide design (primers and TaqMan probes), in silico and in vitro evaluation of design effectiveness, optimization of the physicochemical conditions of the amplification reaction, and result analysis based on experimental designs and acceptance criteria. Furthermore, recommendations are provided for the analytical and clinical validation of the intended use. Our approach is cost-effective, particularly during the design and optimization phases. We primarily used open-source bioinformatics software and tools for in silico evaluations for the test design. Subsequently, the process was manually optimized using a CFX96 Dx analyzer, whose technical specifications and performance are homologous to that of the final platform (Panther Fusion^®). Unlike Panther Fusion^®, the CFX96 Dx does not require excess volumes of reagents, samples, and evaluation materials (dead volume) to accommodate potential robotic handling-associated imprecisions. The utilization of the CFX96 Dx analyzer represents a strategic approach to enhancing the efficiency of resources and the optimization of time during LDT optimization.

Our goal is to understand how hematopoietic stem cell precursors emerge from vessels and to visualize their settling in developmental and more definitive niches that will persist in the adult. For this, we use as a biological model the zebrafish, which offers invaluable advantages owing to its transparency and small size, allowing high-resolution imaging and investigations of the entire animal. In vertebrate species, precursors of hematopoietic stem cells emerge from arterial vessels, mainly from the ventral side of the dorsal aorta. From there, they can either reside in the underlying vascular niche and/or pass through the vein to enter the blood circulation and conquer the caudal hematopoietic tissue, a functional equivalent to the fetal liver in mammals. Here, we provide experimental details of a protocol we have recently optimized to identify, based on mRNA in situ hybridization, precursors of hematopoietic stem cells while still embedded in the aortic wall (at the embryonic stage) as well as when they reside in specific niches a few days after emergence (at the early larval stage). Our experimental approach uses RNAscope technology, which allows combining high-sensitivity mRNA detection with high-resolution fluorescence confocal imaging to achieve spatial transcriptomics. Importantly, the small size of the probes allows better penetration inside tissues, which is a significant improvement in comparison to long mRNA probes; this is an invaluable advantage for reaching deeply embedded niches such as the ones of the pronephros region in the larva and, in addition, provides an increased signal-to-noise ratio.

Fluorescent protein biosensors (FPBs) that turn on—go from dark to bright upon binding their ligands—enable the detection of targets in living cells with high sensitivity and spatial localization. Several approaches exist for creating turn-on FPBs, most notably the method that gave rise to the GCaMP family of genetically encoded calcium indicators. However, it remains challenging to modify these sensors to recognize new ligands. We recently developed adaptable turn-on maturation (ATOM) biosensors, in which target recognition by a small binding domain triggers chromophore maturation in the fluorescent protein to which it is attached. ATOM sensors are advantageous because they are generalizable (by virtue of the monobody and nanobody binding domains) and modular (binding domains and fluorescent proteins of various colors can be mixed and matched for multiplexed imaging), capable of detecting endogenously expressed proteins, and able to function in subcellular compartments including the cytoplasm, nucleus, endoplasmic reticulum, and mitochondria. The protocols herein detail how to design, clone, and screen new ATOM sensors for detecting targets of choice. The starting materials are the genes encoding for a monobody or nanobody and for a cyan, yellow, or red fluorescent protein. We also present general guidelines for creating ATOM sensors using binding domains other than nanobodies and monobodies.

The early detection of meningitis pathogens—including Haemophilus influenzae, Neisseria meningitidis, Streptococcus pneumoniae, and Klebsiella pneumoniae—through point-of-care (POC) systems is essential for mitigating the risk of neurological damage, enhancing patient outcomes, and facilitating prompt clinical decision-making. Nucleic acid amplification testing (NAAT) is a promising tool for improving the diagnosis process of bacterial pathogens associated with brain inflammation. This is due to its high sensitivity, rapidity, and compatibility with portable diagnostic platforms, making it particularly suitable for POC applications. This protocol introduces an innovative diagnostic approach designed to function effectively without the need for advanced laboratory equipment. By leveraging dual-priming isothermal amplification (DAMP), the assay uses custom internal primers to enhance specificity and minimize false results. Brilliant Green is used in this assay for fluorescence detection due to its availability, high fluorescence level, and optimal sample-to-background (S/B) ratio. The assay demonstrated excellent specificity, absence of false positives, sensitivity comparable to loop-mediated isothermal amplification (LAMP), and a high S/B ratio.

Traditional approaches for the detection and differentiation of Bacillus cereus group species often face challenges due to the complexity of genetic discrimination between species. In this protocol, we propose a simple and straightforward assay based on the detected unamplified bacterial 16S rRNA by DNA nanomachine (DNM). The assay incorporates a universal fluorescent reporter and four DNA binding fragments, three of which are responsible for “opening up” the folded rRNA while the fourth strand is responsible for detecting single nucleotide variation (SNV) with high selectivity. The binding of the DNM to 16S rRNA results in the formation of the 10-23 DNAzyme catalytic core that cleaves the fluorescent reporter and produces a signal, which is amplified over time due to catalytic turnover. The developed biplex assay enables the detection of B. thuringiensis 16S rRNA and B. mycoides at fluorescein and Cy5 channels, respectively. The protocol offers two detection options: one utilizing extracted total RNA and the other involving crude cell lysate. The latter enables a fast and straightforward detection after 1.5 h with a hands-on time of ~15 min. The new protocol may simplify the analysis of biological RNA samples and might be useful for environmental monitoring as a simple and inexpensive alternative to amplification-based nucleic acid analysis.

This manuscript details two modified protocols for the isolation of long-stranded or high molecular weight (HMW) DNA from Magnaporthaceae (Ascomycota) fungal mycelium intended for whole genome sequencing. The Cytiva Nucleon PhytoPure and the Macherey-Nagel NucleoBond HMW DNA kits were selected because the former requires lower amounts of starting material and the latter utilizes gentler methods to maximize DNA length, albeit at a higher requirement for input material. The Cytiva Nucleon PhytoPure kit successfully recovered HMW DNA for half of our fungal species by increasing the amount of RNase A treatment and adding in a proteinase K treatment. To reduce the impact of pigmentation development, which occurs toward later stages of culturing, extractions were run in quadruplicate to increase overall DNA concentration. We also adapted the Macherey-Nagel NucleoBond HMW DNA kit for high-quality HMW DNA by grinding the sample to a fine powder, overnight lysis, and splitting the sample before washing the precipitated DNA. For both kits, precipitated DNA was spooled out pre-washing, ensuring a higher percentage of high-integrity long strands. The Macherey-Nagel protocol offers advantages over the first through the utilization of gravity columns that provide gentler treatment, yielding >50% of high-purity DNA strands exceeding 40 kbp. The limitation of this method is the requirement for a large quantity of starting material (1 g). By triaging samples based on the rate of growth relative to the accumulation of secondary metabolites, our methodologies hold promise for yielding reliable and high-quality HMW DNA from a variety of fungal samples, improving sequencing outcomes.

Zebrafish genetic mutants have emerged as a valuable model system for studying various aspects of disease and developmental biology. Mutant zebrafish embryos are generally identified based on phenotypic defects at later developmental stages, making it difficult to investigate underlying molecular mechanisms at earlier stages. This protocol presents a PCR-based genotyping method that enables the identification of wild-type, heterozygous, and homozygous zebrafish genetic mutants at any developmental stage, even when they are phenotypically indistinguishable. The approach involves the amplification of specific genomic regions using carefully designed primers, followed by gel electrophoresis. This genotyping method facilitates the investigation of the molecular mechanisms driving phenotypic defects that are observed at later timepoints. This protocol allows researchers to perform analyses such as immunofluorescence, RT-PCR, RNA sequencing, and other molecular experiments on early developmental stages of mutants. The availability of this protocol expands the utility of zebrafish genetic mutants for elucidating the molecular underpinnings of various biological processes throughout development.

Antimicrobial peptides are effective agents against various pathogens, often targeting essential processes like protein translation to exert their antimicrobial effects. Traditional methods such as puromycin labeling have been extensively used to measure protein synthesis in mammalian and yeast systems; however, protocols tailored for plant pathogenic filamentous fungi, particularly those investigating translation inhibition by antifungal peptides, are lacking. This protocol adapts puromycin labeling to quantify translation inhibition in Botrytis cinerea germlings treated with antifungal peptides. Optimizing the method specifically for fungal germlings provides a precise tool to investigate peptide effects on fungal protein synthesis, advancing our understanding of translation dynamics during pathogen–host interactions in filamentous fungi.

Protein O-GlcNAcylation is a prevalent and dynamic post-translational modification that targets a multitude of nuclear and cytoplasmic proteins. Through the modification of diverse substrates, O-GlcNAcylation plays a pivotal role in essential cellular processes, including transcription, translation, and protein homeostasis. Dysregulation of O-GlcNAc homeostasis has been implicated in a variety of diseases, including cardiovascular diseases, cancer, and neurodegenerative diseases. Studying O-GlcNAcylated proteins in different tissues is crucial to understanding the pathogenesis of these diseases. However, identifying phenotype-relevant candidate substrates in a tissue-specific manner remains unfeasible. We developed a novel tool for the analysis of O-GlcNAcylated proteins, combining a catalytically inactive CpOGA mutant CpOGA^CD and TurboID proximity labeling technology. This tool converts O-GlcNAc modifications into biotin labeling, enabling the enrichment and mass spectrometry (MS) identification of O-GlcNAcylated proteins in specific tissues. Meanwhile, TurboID-CpOGA^DM, which carries two point mutations that inactivate both its catalytic and binding activities toward O-GlcNAc modification, was used as a control to differentiate O-GlcNAc-independent protein–protein interactions. We have successfully used TurboID-CpOGA^CD/DM (TurboID-CpOGA^M) to enrich O-GlcNAc proteins in Drosophila combining the UAS/Gal4 system. Our protocol provides a comprehensive workflow for tissue-specific enrichment of candidate O-GlcNAcylated substrates and offers a valuable tool for dissecting tissue-specific O-GlcNAcylation functions in Drosophila.

Transfer RNAs (tRNAs), the essential adapter molecules in protein translation, undergo various post-transcriptional modifications. These modifications play critical roles in regulating tRNA folding, stability, and codon–anticodon interactions, depending on the modified position. Methods for detecting modified nucleosides in tRNAs include isotopic labeling combined with chromatography, antibody-based techniques, mass spectrometry, and high-throughput sequencing. Among these, high-performance liquid chromatography (HPLC) has been a cornerstone technique for analyzing modified nucleosides for decades. In this protocol, we provide a detailed, streamlined approach to purify and digest tRNAs from yeast cells and analyze the resulting nucleosides using HPLC. By assessing UV absorbance spectra and retention times, modified nucleosides can be reliably quantified with high accuracy. This method offers a simple, fast, and accessible alternative for studying tRNA modifications, especially when advanced technologies are unavailable.