微生物学

Microbial life cycles are often reconstructed theoretically from fragmentary pieces of evidence. Protocols for the direct and continuous observation of entire microbial life cycles, including sexual reproduction, are scarce, which limits the study of cellular transitions between different life cycle stages and prevents the visualization of cryptic stages. Although sequence-based techniques, such as -omics approaches, can reconstruct cellular transitions at the genetic and biochemical level, these methods are destructive and do not recover information from the same living cell over time. This protocol provides a solution to directly and continuously observe microbial life cycles, including sexual reproduction, by using microfluidics manipulations that expose single cells to nutritional stimuli and selective pressures. As proof of principle, we triggered a life cycle sequence transition in the model yeast Saccharomyces cerevisiae, starting with an arrest of proliferation in an ancestor cell followed by induction of meiosis through starvation, selection of sexually reproducing cells through exposure to a drug cocktail, germination of haploid spores, and mating of haploid individuals, creating a new descendant generation. This protocol offers the possibility to directly compare molecular and cellular behavior across life cycle stages and across sexually reproducing generations.

Lipid droplets have emerged as dynamic organelles involved in diverse cellular processes beyond simple lipid storage. In plants and cyanobacteria, growing evidence highlights their importance in stress adaptation and signaling, yet methods to study their structure and purity remain limited. Traditionally, in situ transmission electron microscopy (TEM) has been used to visualize lipid droplets within intact cells. While powerful, this approach cannot easily evaluate isolated lipid droplets or confirm their purity. In this protocol, we describe a rapid method for preparing and visualizing cyanoglobule lipid droplets isolated from cyanobacteria. The isolated droplets are directly processed for TEM using negative staining with uranyl acetate, providing a straightforward and efficient workflow. The procedure can be applied broadly to lipid droplets from diverse organisms, independent of species or cellular origin. This protocol offers a simple, fast, and widely applicable approach to assessing lipid droplets, expanding the toolkit for researchers studying their structure and function.

Quantitative analysis of biological membrane morphology is essential for understanding fundamental cellular processes such as organelle biogenesis and remodeling. While manual annotation has been the standard for complex structures, it is laborious and subjective, and conventional automated methods often fail to accurately delineate overlapping objects in 2D projected microscopy images. This protocol provides a complete, step-by-step workflow for the quantitative analysis of overlapping prospore membranes (PSMs) in sporulating yeast. The procedure details the synchronous induction of sporulation, acquisition of 3D fluorescence images and their conversion to 2D maximum intensity projections (MIPs), and the generation of a custom-annotated dataset using a semi-automated pipeline. Finally, it outlines the training and application of our mask R-CNN-based model, DeMemSeg, for high-fidelity instance segmentation and the subsequent extraction of morphological parameters. The primary advantage of this protocol is its ability to enable accurate and reproducible segmentation of individual, overlapping membrane structures from widely used 2D MIP images. This framework offers an objective, efficient, and scalable solution for the detailed quantitative analysis of complex membrane morphologies.

Genome-walking protocols have been extensively used to clone unknown genomic sequences next to known DNAs. Existing genome-walking protocols need further improvement in methodological specificity or operation. Here, we describe a novel genome-walking protocol based on fusion primer–driven racket PCR (FPR-PCR). FPR-PCR involves four sequence-specific oligos (SSO), SSO1, SSO2, SSO3, and SSO4, which are sequentially chosen from known DNA in the direction 5’→3’. The fusion primer, mediating primary FPR-PCR, is generated by attaching SSO3 to the 5’ end of SSO1. The SSO3 encourages the target DNA of primary PCR to form a racket-like structure by mediating intra-strand annealing. SSO2 and SSO4 are directly used as sequence-specific primers (SSP) in secondary FPR-PCR, which selectively amplifies this racket-like DNA. This protocol was verified by cloning several unknown genomic sequences. Compared to traditional PCRs, FPR-PCR offers the advantages of higher specificity and fewer rounds, primarily attributed to the omission of arbitrary walking primers typically required in traditional methods.

Xenobiotics, including environmental pollutants such as bisphenols, phthalates, and parabens, are widely present in food, cosmetics, packaging, and water. These compounds can reach the gastrointestinal tract and interact with the gut microbiota (GM), a complex microbial community that plays a key role in host immunity, metabolism, and barrier function. The GM engages in bidirectional communication with the host via the production of bioactive metabolites, including short-chain fatty acids, neurotransmitter precursors, and bile acid derivatives. Dysbiosis induced by xenobiotics can disrupt microbial metabolite production, impair gut barrier integrity, and contribute to the development of systemic disorders affecting distant organs such as the liver or brain. On the other hand, the GM can biotransform xenobiotics into metabolites with altered bioactivity or toxicity. In vitro models of the human GM offer a valuable tool to complement population-based and in vivo studies, enabling controlled investigation of causative effects and underlying mechanisms. Here, we present an optimized protocol for the collection, cryopreservation, and cultivation of human GM under strictly anaerobic conditions for toxicomicrobiomics applications. The method allows the assessment of xenobiotic–GM interactions in a cost-effective and ethically sustainable way. It is compatible with a wide range of downstream applications, including 16S rDNA sequencing, metabolomics, and endocrine activity assays. The protocol has been optimized to minimize oxygen exposure to less than 2 min, ensuring the viability of obligate anaerobes that dominate the gut ecosystem. This approach facilitates reproducible, mechanistic studies on the impact of environmental xenobiotics on human GM.

The rhizosphere, a 2–10 mm region surrounding the root surface, is colonized by numerous microorganisms, known as the rhizosphere microbiome. These microorganisms interact with each other, leading to emergent properties that affect plant fitness. Mapping these interactions is crucial to understanding microbial ecology in the rhizosphere and predicting and manipulating plant health. However, current methods do not capture the chemistry of the rhizosphere environment, and common plant–microbe interaction study setups do not map bacterial interactions in this niche. Additionally, studying bacterial interactions may require the creation of transgenic bacterial lines with markers for antibiotic resistance/fluorescent probes and even isotope labeling. Here, we describe a protocol for both in silico prediction and in vitro validation of bacterial interactions that closely recapitulate the major chemical constituents of the rhizosphere environment using a widely used Murashige & Skoog (MS)-based gnotobiotic plant growth system. We use the auto-fluorescent Pseudomonas, abundantly found in the rhizosphere, to estimate their interactions with other strains, thereby avoiding the need for the creation of transgenic bacterial strains. By combining artificial root exudate medium, plant cultivation medium, and a synthetic bacterial community (SynCom), we first simulate their interactions using genome-scale metabolic models (GSMMs) and then validate these interactions in vitro, using growth assays. We show that the GSMM-predicted interaction scores correlate moderately, yet significantly, with their in vitro validation. Given the complexity of interactions among rhizosphere microbiome members, this reproducible and efficient protocol will allow confident mapping of interactions of fluorescent Pseudomonas with other bacterial strains within the rhizosphere microbiome.

Sheath blight, caused by Rhizoctonia solani, is a major fungal disease of rice that leads to significant yield losses globally. Conventional inoculation methods often fail to achieve consistent and uniform infection, limiting their applicability in antifungal screening studies. This protocol describes a reliable in planta inoculation method for R. solani using mature sclerotia placed at the internodal region of tillering-stage rice seedlings. The procedure includes step-by-step instructions for seed germination, seedling preparation, pathogen culture, artificial inoculation, and post-infection application of antifungal treatments, including botanical compounds such as Ocimum gratissimum essential oil and thymol. Lesion development is monitored and quantified over time, and data are analyzed statistically to evaluate treatment efficacy. The protocol is optimized for reproducibility, scalability, and compatibility with sustainable disease management approaches. It provides a robust platform for evaluating antifungal agents in a biologically relevant and controlled environment.

The RNA-guided Cas enzyme specifically cuts chromosomes and introduces a targeted double-strand break, facilitating multiple kinds of genome editing, including gene deletion, insertion, and replacement. Caulobacter crescentus and its relatives, such as Agrobacterium fabrum and Sinorhizobium meliloti, have been widely studied for industrial, agricultural, and biomedical applications; however, their genetic manipulations are usually characterized as time-consuming and labor-intensive. C. crescentus and its relatives are known to be CRISPR/Cas-recalcitrant organisms due to intrinsic limitations of SpCas9 expression and possible CRISPR escapes. By fusing a reporting gene to the C terminus of SpCas9M and precisely manipulating the expression of SpCas9M, we developed a CRISPR/SpCas9M-reporting system and achieved efficient genome editing in C. crescentus and relatives. Here, we describe a protocol for rapid, marker-less, and convenient gene deletion by using the CRISPR/SpCas9M-reporting system in C. crescentus, as an example.

Candida albicans is the pathogenic fungus that most frequently causes infections in humans. It is part of the microbiota commonly found in the skin, gastrointestinal tract, and vaginal mucosa. However, certain conditions, including immunosuppression, excessive use of antibiotics, hormonal changes, the use of medical devices in patients, and individual nutritional status, promote the development of opportunistic infections caused by this fungus. One of the main fungal structures interacting with the host is the cell wall, which is principally composed of chitin, glucan, and proteins. The cell wall plays key functions for the cell, such as osmotic protection; it is also responsible for cellular shape and acts as a signaling hub in response to environmental changes. Cell wall proteins participate in diverse cellular functions, such as attachment to surfaces and cell wall structure; some possess catalytic or transport activities. In this protocol, we show the methodology for isolating cell wall proteins covalently linked or not to cell wall components that can be previously labeled with [¹⁴C]-L-lysine by the action of the fungal transglutaminase localized in the cell wall. We use an extraction method by mechanical cell disruption and washing with 2 M NaCl, whose ionic strength eliminates contaminating proteins from other organelles, through subsequent serial treatments with SDS, chitinase, and zymolyase.

Oomycetes are a predominantly plant-pathogenic group of organisms often considered and managed as fungi; however, due to their evolutionary divergence from true fungi, many conventional fungicides are ineffective against them. Their unique physiological characteristics make them challenging to work with, highlighting the need for a standardized and reproducible procedure for anti-oomycete assays. Previous studies describe methods to obtain sporulation forms in the laboratory, but there remains a disconnect between spore production and the subsequent screening process for potential biological pesticides based on microbial organic extracts. This protocol bridges that gap by providing a complete and reliable workflow from spore production to screening. In this study, we present an efficient in vitro protocol to identify microbial extracts with activity against Phytophthora capsici and Pythium ultimum. The protocol includes a method for obtaining zoospores of P. capsici and oospores of P. ultimum, followed by a simple and rapid screening assay to detect microbial extracts that inhibit the growth of these pathogens. The extracts are dispensed onto plates in two concentrations and allowed to dry. This facilitates pauses in the protocol and allows for storage of the plates until the biological material is ready for the assay. The protocol’s effectiveness has been validated with these two oomycetes, resulting in the identification of active extracts in both cases. Moreover, it can be adapted to other pathogens.