微生物学

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.

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.

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.

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.

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.

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.

Cyanobacteria have been widely used as model organisms in photobiochemical research and have recently been exploited as hosts in numerous pilot studies to produce valuable biochemicals via genetic and metabolic modifications. Analyzing cellular RNA is a suitable method for studying genetic changes in cells. Several methods have previously been reported for cyanobacterial RNA extraction. However, the majority of these methods rely heavily on phenol and chloroform, which are hazardous. Additionally, these methods are time-consuming and difficult to perform. Using Synechocystis sp. PCC 6803 as a model, this study developed a novel method for extracting total ribonucleic acid (RNA) using standard centrifugation techniques and laboratory chemicals such as citric acid, ethylenediaminetetraacetic acid, sodium dodecyl sulfate, sodium chloride, and tri-sodium citrate dihydrate to extract RNA from cyanobacterial cells. This method does not necessitate the use of hazardous chemicals, especially phenol and chloroform. Furthermore, it is cost-effective since it does not require expensive chemicals. The results of the quantification, purity, and integrity checks show the effectiveness of this method for extracting good-quality RNA. Furthermore, RT-qPCR results demonstrate that the quality of the extracted RNA is suitable for downstream applications.

Mal de Río Cuarto disease, caused by a Fijivirus, is a major constraint for maize production in Argentina. The traditional evaluation of resistant hybrids is limited by the low efficiency of natural virus transmission and the lack of standardized field inoculation methods. We developed a protocol that combines laboratory mass-rearing of the planthopper vector Delphacodes kuscheli with a controlled field transmission system. The method involves the synchronized production of large insect populations, acquisition of viruliferous vectors under controlled conditions, and their safe transport to the field using specialized containers. Transmission is achieved through individual cages placed on maize seedlings, ensuring high inoculation pressure under field-like conditions. This protocol enables reliable and reproducible virus transmission, facilitating large-scale screening of maize hybrids and other cereals. Its main advantages are the high throughput of vector production, improved transmission efficiency, and adaptability to diverse experimental designs.

Single-cell RNA sequencing (scRNA-seq) is a powerful technique for exploring cellular heterogeneity and host–pathogen interactions. This protocol details the Zika virus (ZIKV)-targeted scRNA-seq workflow for preparing high-quality single-cell suspensions from the whole brain tissues of neonatal mice, high-quality single-cell sorting, cDNA reverse transcription, amplification, ZIKV enrichment and host transcriptome library preparation, and sequencing dataset integration in downstream analysis to complete the quantification of ZIKV RNA in individual cells.

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.