Microbiology

Pathogen gene expression from host samples is often challenging to study due to low signal and high host RNA background. PCR probes have been recently used to hybridize and extract bacterial sequences from next-generation sequencing (NGS) libraries generated from in vitro and animal models of infection; however, these strategies require purchasing commercially synthesized probes that often do not capture the entire transcriptome. Transcript-capture sequencing is a novel capture approach for extracting RNA of a target bacterial species from samples in which there is substantial contamination by the host or other microbes. Biotinylated 150-base-pair DNA probes are generated in-house from bacterial DNA spanning the entire bacterial genome. Probes are hybridized to the cDNA of NGS sequencing libraries prepared from host samples to capture and enrich for bacterial-specific RNA reads before sequencing. This method results in a >200-fold increase in bacterial RNA reads from infected host samples (including in vitro, animal, and human samples) and generates complete bacterial transcriptomes with high gene coverage (>80%). Use of this protocol on infected host samples reveals a snapshot of bacterial activity during disease that may improve understanding of the physiological state of pathogens within their hosts.

The genomes of RNA viruses can fold into dynamic structures that regulate their own infection and immune evasion processes. Proximity ligation methods (e.g., SPLASH) enable genome-wide interaction mapping but lack specificity when dealing with low-abundance targets in complex samples. Here, we describe HiCapR, a protocol integrating in vivo psoralen crosslinking, RNA fragmentation, proximity ligation, and hybridization capture to specifically enrich viral RNA–RNA interactions. Captured libraries are sequenced, and chimeric reads are analyzed via a customized computational pipeline to generate constrained secondary structures. HiCapR generates high-resolution RNA interaction maps for viral genomes. We applied it to resolve the in vivo structure of the complete HIV-1 RNA genome, identifying functional domains, homodimers, and long-range interactions. The protocol's robustness has been previously validated on the SARS-CoV-2 genome. HiCapR combines proximity ligation with targeted enrichment, providing an efficient and specific tool for studying RNA architecture in viruses, with broad applications in virology and antiviral development.

The CRISPR/Cas12a system has revolutionized molecular diagnostics; however, conventional Cas12a-based methods for RNA detection typically require transcription and pre-amplification steps. Our group has recently developed a diagnostic technique known as the SCas12a assay, which combines Cas12a with a split crRNA, achieving amplification-free detection of miRNA. However, this method still encounters challenges in accurately quantifying long RNA molecules with complex secondary structures. Here, we report an enhanced version termed SCas12aV2 (split-crRNA Cas12a version 2 system), which enables direct detection of RNA molecules without sequence limitation while demonstrating high specificity in single-nucleotide polymorphism (SNP) applications. We describe the general procedure for preparing the SCas12a system and its application in detecting RNA targets from clinical samples.

Chemical phenotyping is a fundamental technique to study the metabolic properties or chemical sensitivities of bacteria. Traditional methods such as dilution methods, discs, or gradient diffusion assays are labor-intensive, often have high material requirements, and are limited in scalability. High-throughput cultivation approaches based on 96-well plates scale efficiently to large numbers of samples. A stacker, when coupled with a plate reader system (often already available in most laboratories), greatly enhances assay scalability and robustness. Here, we describe a customized high-throughput, flexible, scalable, robust, and affordable method for the chemical phenotyping of bacteria. This liquid culture–based growth system allows screening many bacteria in parallel and in a replicated manner for their tolerance to various chemicals, including specialized metabolites of plants, antibiotics, or pesticides. Compared to commercial solutions, our approach offers high flexibility in experimental conditions while keeping costs for consumables low.

Vulvovaginal candidiasis (VVC), also known as vaginal thrush, is an infection of the vulvovaginal mucosa caused by fungi of the Candida genus. Particularly for patients suffering from recurrent infection, the disease has a significant impact on their quality of life. The still unknown aspects of disease pathogenesis, as well as factors driving the development of infections and recurrence, represent a challenge for both clinical practitioners and patients. Mouse models and patient studies have suggested important roles of the microbiome, deployment of fungal pathogenicity mechanisms in the vagina, and dysregulated immune responses for VVC pathology. Dissecting their individual contributions can reveal specific processes associated with infection and may inspire novel therapeutic strategies. Epithelial in vitro infection models have been playing a key role in dissecting a crucial interaction during VVC, the invasion and infection of the vaginal mucosa. They have been instrumental in characterizing candidalysin as a fungal toxin that damages epithelial cells and elicits initial inflammatory responses to catalyze downstream inflammation. Moreover, they have also revealed potential protective immune pathways. Such a standardized epithelial cell infection model offers high versatility and compatibility with different downstream assays to link epithelial responses with other processes during VVC. This protocol describes a general A-431 vulvovaginal epithelial cell–Candida infection model in detail and provides several adaptations, such as live-cell imaging and mRNA silencing, as well as possible follow-up readouts, like the quantification of cytokine release, cytotoxicity, and neutrophil recruitment to study diverse processes relevant to VVC research.

Pseudoalteromonas haloplanktis TAC125 is a psychrophilic marine bacterium widely used to study cold adaptation and increasingly exploited as a non-conventional platform for biotechnological applications. The strain harbors the endogenous megaplasmid pMEGA (64.7 kb), whose presence may limit its exploitation as a cell factory, making its elimination advantageous to strain engineering. Traditional plasmid-curing approaches based on chemical and physical agents are often inefficient and unsuitable for stable endogenous replicons, such as pMEGA. Here, we describe a targeted protocol for pMEGA curing in P. haloplanktis TAC125 that combines homologous recombination with paired-termini antisense RNA (PTasRNA) gene silencing. First, a selectable marker cassette is inserted into pMEGA by homologous recombination using a suicide vector, enabling selective discrimination between plasmid-positive and plasmid-cured bacteria. Next, PTasRNA gene silencing technology is applied to target a gene essential for the replication of pMEGA, thereby transiently interfering with its replication and promoting its loss. This approach provides a specific method to cure a highly stable endogenous megaplasmid in a psychrophilic non-conventional bacterium, enabling improved functional studies and strain optimization, establishing a broadly applicable framework for targeted curing across diverse bacterial systems.

3-nitro-tyrosine (nitroTyr) is one of numerous oxidative protein modifications implicated in diseases such as cardiovascular disease, cancer, and amyotrophic lateral sclerosis (ALS). Because of this, the ability to site-specifically encode nitroTyr into recombinant proteins is a powerful approach for studying these disease pathways. However, producing proteins with defined nitration sites is technically challenging due to the limitations of traditional chemical nitration via peroxynitrite, which lacks residue and site-specificity. Genetic code expansion (GCE) offers a solution by enabling precise incorporation of nitroTyr at designated TAG codons using engineered aminoacyl-tRNA synthetase/tRNA pairs from Methanocaldococcus jannaschii and Methanomethylophilus alvus. This protocol provides a reliable, optimized workflow for incorporating nitroTyr into proteins in E. coli using GCE. It guides users through key considerations in selecting cell lines, media conditions, and GCE systems to minimize off-target effects such as release factor 1 competition, near-cognate suppression, and chemical reduction of nitroTyr. The method is demonstrated using wild-type and TAG-containing superfolder GFP but is broadly applicable to other proteins of interest.

Engineering of microbial cells, including E. coli, is essential in prototyping genetic designs used in numerous applications throughout synthetic biology. While many advanced genome editing tools, such as CRISPR-based tools, offer new capabilities with genetically recalcitrant organisms, these tools often do not offer an immediate advantage in readily manipulated microbes, such as E. coli, especially when scarless modifications are not critical. We describe a comprehensive recombineering tutorial that we commonly use for multiplex engineering of E. coli using antibiotic markers. We leverage a group of 15 antibiotic resistance cassettes, most of which can be readily included when designing double-stranded DNA donors intended for recombineering and purchased from several vendors. Using these methods, 10–15 defined modifications to a single host strain can be achieved in less than three weeks, using two-day editing cycles. We discuss sequences and protocols as well as the optimal design of genetic modifications and the associated DNA.

Protein–protein interactions (PPIs) govern nearly all aspects of cellular physiology, yet identifying these interactions under native conditions remains challenging. Here, we present TIE-UP-SIN (targeted interactome experiment for unknown proteins by stable isotope normalization), a robust method for in vivo identification and quantification of PPIs in bacterial systems. The protocol combines metabolic labeling with ¹⁵N isotopes, reversible formaldehyde crosslinking, affinity purification, and quantitative mass spectrometry. TIE-UP-SIN preserves transient or weak interactions during purification and quantifies interaction partners using internal light/heavy peptide ratios, reducing experimental variability. The method employs a triple-sample design to distinguish specific from nonspecific interactors and can be adapted to various bacterial species and affinity tags. Data analysis is streamlined through a user-friendly web application (https://shiny-fungene.biologie.uni-greifswald.de/TIE_UP_SIN_app) that automates statistical analysis, normalization, and visualization, requiring no programming expertise. The entire workflow from cell culture to mass spectrometry data acquisition takes approximately 4–5 days, with data analysis completed in 1–2 days using the web application.

The rising global incidence of pancreatitis, pancreatic cancer, and diabetes has increased the need for efficient in vivo gene manipulation approaches to study the pancreas and develop new therapies. Although transgenic mouse models are widely used, they are time-consuming and costly to generate and maintain. Systemic viral delivery methods offer greater flexibility but often lack pancreatic specificity and require high viral doses. Here, we describe a streamlined protocol for intrapancreatic ductal delivery of adeno-associated viruses (AAVs) for targeted gene delivery. Our protocol requires standard surgical equipment and can be implemented in most laboratories. Specifically, we adopted a clamping strategy at the hepatopancreatic duct near the liver, as well as beneath the major duodenal papilla at the duodenum. This strategy exposes the duodenal papilla, facilitating viral delivery, preventing backflow, and enabling efficient pancreatic transduction at lower viral doses. Overall, this method provides a fast, simple, and effective approach for pancreas-targeted gene manipulation, facilitating preclinical studies of pancreatic biology and disease.