Microbiology

Isogenic populations of Saccharomyces cerevisiae exhibit significant proliferative heterogeneity, with individual cells within a clonal culture displaying divergent growth rates and metabolic states. Investigating the origins of this variation requires a method to reconstruct the individual histories of cells within the population. This protocol describes a method for single-cell microencapsulation in alginate microspheres to create a physically stable, traceable, three-dimensional genealogical environment. By utilizing the alginate matrix to prevent daughter cell migration, the replicative history of a founder cell can be mathematically reconstructed. This is achieved by correlating the total cell count (N) within a developed microcolony with the total number of accumulated bud scars (n) visualized via confocal microscopy.

Bacterial cellulose (BC) is a renewable biopolymer valued for its exceptional purity, biocompatibility, and mechanical strength, with broad applications in biomedicine and sustainable materials. However, achieving reproducible BC production and downstream processing remains a major challenge. Inoculum preparation is particularly difficult to standardize because cellulose-producing strains form pellicles that sequester cells, making optical density measurements unreliable. In addition, recovery and drying procedures can alter fiber accessibility, and enzymatic hydrolysis conditions are often inconsistently defined and lack proper enzyme activity assessment. These issues contribute to substantial variability in BC-derived nanoparticle yields. This protocol describes the production of BC from Komagataeibacter xylinus DSMZ 6513, including culture medium preparation, inoculum generation, and scaling up under static cultivation conditions. It further details BC pellicle purification using NaOH, followed by pulping, freeze-drying, and milling to ensure material stability during storage and use. BC hydrolysis is performed with commercially available cellulase from Trichoderma reesei, with enzyme activity quantified prior to each reaction to ensure reproducibility. This standardized approach enables the reproducible production of bacterial cellulose nanoparticles (BCNPs). The protocol also includes minimal morphological characterization methods. By standardizing culture, recovery, and hydrolysis steps, the workflow reduces experimental variability and improves comparability across laboratories. Overall, it provides an accessible and reproducible method for generating BC and BCNPs of consistent quality without the need for specialized instrumentation.

Mycoparasitism is an important mechanism of fungal antagonism in which one fungus parasitizes another. This type of interaction plays a major role in the biocontrol activity of Trichoderma spp. against phytopathogenic fungi. Detailed visualization of these interactions is essential for understanding the structural mechanisms involved in fungal antagonism, including hyphal attachment, coiling, penetration, and cellular distortion. Scanning electron microscopy (SEM) is widely used for structural examination of fungal interactions; however, conventional preparation methods such as filter paper systems, membrane overlays, and agar block techniques often result in structural distortion, fragile sample handling, and difficulty in locating defined interaction zones. Here, we describe a modified slide-embedded technique for SEM visualization of mycoparasitic interactions between filamentous fungi. The protocol is adapted from previously reported slide culture approaches and involves embedding pre-cut sterile glass slide fragments directly into potato dextrose agar (PDA), followed by sequential inoculation of Fusarium sp. and Trichoderma viride. Fungal interactions occurring directly on the glass surface are subsequently subjected to fixation with 2.5% glutaraldehyde, graded ethanol dehydration, sputter coating, and SEM observation. Compared with conventional methods, the present approach provides improved handling stability, better preservation of native hyphal architecture, reduced deformation during processing, and easier localization of interaction zones during microscopy. The protocol also enables clear visualization of early antagonistic events such as hyphal coiling, penetration, and surface colonization. Due to its simplicity, reproducibility, and minimal technical complexity, this method serves as a practical and efficient approach for SEM-based investigation of fungal–fungal interactions and can be readily adapted for studying diverse mycoparasitic systems.

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.

Cyclic di-AMP is a bacterial second messenger nucleotide required for the regulation of numerous cellular functions, including potassium and osmolyte homeostasis, DNA repair, cell wall integrity, central metabolism, and stress adaptation. This second messenger is synthesized from two ATP molecules by diadenylate cyclases (DAC) and degraded by cytoplasmic and surface-associated phosphodiesterases (PDE) to phosphoadenylyl adenosine (5′ pApA), adenosine monophosphate (AMP), and, in some instances, adenosine and inorganic phosphate (Pi). Levels of c-di-AMP in bacteria can be determined using different methods, including liquid chromatography–mass spectrometry (LC-MS/MS), enzyme-linked immunosorbent assay (ELISA), and luminescent and fluorescent biosensors. Thin-layer chromatography (TLC) is another method routinely used to monitor c-di-AMP synthesis and degradation by purified DAC and PDE enzymes and is particularly useful for monitoring c-di-AMP degradation products. Here, we devised a TLC-based method to monitor extracellular c-di-AMP stability and degradation by intact bacterial cells using radiolabeled c-di-AMP. We show that bacterial strains of Enterococcus faecalis and Streptococcus agalactiae that possess surface-associated PDEs can rapidly degrade extracellular c-di-AMP. In addition, we demonstrate that this method can be used to indirectly identify alternative enzyme substrates through competition assays. We propose that this TLC-based assay is an efficient method to analyze bacterial-mediated degradation of c-di-AMP and is amenable to testing other radiolabeled nucleotides.

Difficult-to-treat Gram-negative bacteria are a major cause of healthcare-associated infections due to multidrug resistance and limited therapeutic options. The hospital environment plays a central role in the persistence and transmission of infection, making environmental monitoring an essential component of infection prevention and control plans. Standard surface sampling techniques, including swabs, contact plates, and sponges, are widely used for environmental surveillance and are all based on culture-dependent methods. However, these techniques may underestimate the actual level of bacterial contamination since they fail to detect bacteria in the viable but non-culturable (VBNC) state, a reversible physiological condition in which bacterial cells remain viable but do not grow on conventional culture media. An innovative environmental sampling protocol, herein named MORECOVERY, has been developed to improve the detection of VBNC bacteria. The protocol integrates an essential resuscitation step, which improves the recovery of VBNC Gram-negative bacteria by a few orders of magnitude, into the standard swab-based sampling workflow. Following sample collection, swabs are incubated for 24 h at 37 °C in a carbon-free resuscitation buffer before plating, enabling the recovery of VBNC bacterial pathogens that would otherwise remain undetectable. By improving the recovery of VBNC cells, the MORECOVERY protocol allows a more accurate assessment of bacterial contamination of critical surfaces in healthcare settings. Its simplicity and minimal variation from standard workflows facilitate easy implementation in routine environmental monitoring.

Hepatitis E virus (HEV) is a zoonotic pathogen responsible for approximately 20 million infections annually worldwide. The lack of robust cell culture systems and the absence of approved antiviral therapies have hindered HEV research and drug development. A major technical challenge is the rapid loss of viral infectivity during freeze–thaw cycles following virus purification. Here, we describe a simple and reproducible method to preserve HEV infectivity during storage. We systematically evaluated the effects of salt, serum, and sucrose on viral stability under freezing conditions. We identified an optimized buffer containing 2% fetal bovine serum (FBS), 150 mM NaCl, and 7% sucrose, which significantly maintained the infectivity of non-enveloped HEV (nHEV) and quasi-enveloped HEV (eHEV) following freeze–thaw cycles based on immunofluorescence. The buffer also demonstrated good stability across three independent repeat infection experiments. This protocol provides a practical and scalable approach for maintaining HEV infectivity and will facilitate HEV-related virological studies.

RNA sequencing (RNA-seq) has revolutionized transcriptomics, ribosome footprinting, and polysome profiling, providing a wealth of data. Many RNA-based omics typically remove ribosomal RNA (rRNA) or select for messenger RNA (mRNA) prior to sequencing, thereby enriching reads that map to the translationally active part of the transcriptome. Prokaryotic mRNA lacks the 3′ polyadenylated tail, which excludes the use of poly(A)-based selection methods. While commercial rRNA depletion products exist for prokaryotes, their proprietary nature and potential inefficiency with non-model organisms are factors that may limit broad-scale application. To mitigate this issue, we designed DepStep, a consolidated workflow for one-step rRNA depletion using species-specific biotinylated antisense probes for selective hybridization and removal of the target rRNA molecules. As a proof-of-concept, RNA-seq libraries of the psychrophilic gram-negative bacterium Shewanella glacialimarina TZS-4_T were prepared using both DepStep and a commercial rRNA depletion kit for gram-negative bacteria, to which DepStep was benchmarked. DepStep compares favorably to the commercial depletion kit; it removes >98.6% of the rRNA content in the sample, resulting in sequencing libraries where the coding DNA sequence (CDS) reads account for >80% of the total read count. Importantly, DepStep’s cost-per-sample is three times lower than the commercial kit, establishing DepStep as a simple yet cost-effective alternative to commercial solutions.

Understanding microbial survival under extreme planetary conditions is critical for astrobiology and stress biology. Several experimental platforms, including radiation, desiccation, and microgravity, have been used to mimic extraterrestrial environments; however, controlled simulation of high-intensity shock waves has not been used to assess microbial survival. Here, we describe a detailed protocol for shock processing of Saccharomyces cerevisiae using the high-intensity shock tube for astrochemistry (HISTA), which generates high-Mach-number shock waves under inert gas conditions. Yeast cells are drop-casted onto a metal flange, exposed to transient high-pressure shock waves, and recovered for downstream survival and cellular analyses. Shock intensity can be precisely tuned by adjusting driver pressure, diaphragm thickness, and driven gas pressure. This protocol provides a platform to investigate microbial adaptation to shock waves.

Careful dissection of insect gut tissues is essential for microbiome studies to ensure accurate characterization of internal microbial communities and preservation of DNA integrity. Because insect-associated microbiomes are highly sensitive to contamination, effective removal of external microbes prior to dissection is critical to minimize bias in downstream analyses. While ethanol- and bleach-based surface sterilization methods are commonly used, standardized workflows integrating field collection, sterilization, and dissection remain limited. Here, we present a step-by-step protocol for the field collection, surface sterilization, and dissection of gut tissues from the agricultural pest Diaprepes abbreviatus (Coleoptera: Curculionidae), optimized for genomic DNA extraction and microbiome analyses. Using wild-caught specimens, this workflow incorporates a rigorous surface sterilization and dissection strategy that minimizes external contamination while preserving biologically relevant microbial signatures and DNA integrity for downstream microbiome analyses. The protocol provides a standardized framework for insect gut microbiome studies and can be broadly adapted to other wild-caught insect species requiring careful collection, disinfection, and sterile dissection prior to molecular analysis. The protocol integrates field collection and laboratory processing steps into a streamlined workflow that minimizes contamination while preserving tissue integrity for downstream applications.