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.

Cryo-electron microscopy (cryo-EM) is a powerful technique capable of investigating samples in a hydrated state, compared to conventional high-vacuum electron microscopy that requires samples to be completely dry. During the drying process, numerous features and details may be lost due to damage caused by dehydration. Cryo-EM circumvents these problems by cryo-fixing the samples, thereby retaining the intact and original features of hydrated samples. This protocol describes a step-by-step cryo-scanning electron microscopy (cryo-SEM) experimental procedure with Chlorella sorokiniana as the subject. By employing filter paper as the sample substrate, we propose a simple and reliable method for cryo-fixation and freeze-fracture of Chlorella sorokiniana in water suspension. The advantage of using filter paper as a substrate lies in its ability to support a thin film of sample, enabling a cold knife to make a cut effortlessly and produce a clean freeze-fractured surface for SEM investigation. By following the approach described in this protocol, both the internal structure and surface morphology of Chlorella sorokiniana can be easily resolved with high quality. This protocol is highly versatile and can be applied to samples dispersed in water or solvents, including cyanobacterial cells, algal cells, and any kind of sample that can be adsorbed onto filter paper.

Surface proteins of Staphylococcus aureus and other Gram-positive bacteria play essential roles in bacterial colonization and host-microbe interactions. Surface protein precursors containing a YSIRK/GXXS signal peptide are translocated across the septal membrane at mid-cell, anchored to the cell wall peptidoglycan at the cross-wall compartment, and presented on the new hemispheres of the daughter cells following cell division. After several generations of cell division, these surface proteins will eventually cover the entire cell surface. To understand how these proteins travel from the bacterial cytoplasm to the cell surface, we describe a series of immunofluorescence microscopy protocols designed to detect the stepwise subcellular localization of the surface protein precursors: surface display (protocol A), cross-wall localization (protocol B), and cytoplasmic/septal membrane localization (protocol C). Staphylococcal protein A (SpA) is the model protein used in this work. The protocols described here are readily adapted to study the localization of other surface proteins as well as other cytoplasmic or membrane proteins in S. aureus in general. Furthermore, the protocols can be modified and adapted for use in other Gram-positive bacteria.

Graphic abstract:

Tracking the subcellular localization of surface proteins in S. aureus

Detecting live bacteria is an important task for antimicrobial susceptibility testing (AST) in the medical sector and for quality-monitoring in biological industries. Current methods for live-bacteria detection suffer limitations in speed or sensitivity. In a recent paper, we reported that electrical response dynamics in membrane potential enable single-cell rapid detection of live bacteria. The electrical response can be observed within a minute after electrical stimulation. Thus, it has potential in accelerating AST and the monitoring of biological samples. This method also enables experiments for biophysical and microbiological investigations into bacterial electrophysiology. With the hope that more researchers, scientists and engineers will use electrical stimulation for their assays, here we detail each step of the electrical stimulation experiment.

Extracellular vesicles (EVs) are produced by all domains of life including Bacteria, Archaea and Eukarya. EVs are critical for cellular physiology and contain varied cargo: virulence factors, cell wall remodeling enzymes, extracellular matrix components and even nucleic acids and metabolites. While various protocols for isolating EVs have been established for mammalian cells, the field is actively developing tools to study EVs in other organisms. In this protocol we describe our methods to perform density gradient purification of EVs in bacterial cells, allowing for separation of EV subpopulations, followed by protection assays for EV cargo characterization. Furthermore, we devised a protocol which incorporates a fluorescent conjugate of fatty acids into EVs, the first to allow live-cell EV tracking to observe release of EVs, including during infection of mammalian cells by pathogenic bacteria. These protocols are powerful tools for EV researchers as they enable the observation of EV release and the study of the mechanisms of their formation and release.

The most important virulence factor in the Cryptococcus genus is the polysaccharide capsule. This genus includes several species that cause life-threatening invasive disease. An increase in capsule thickness is important during fungal infection. The capsule is usually imaged using India ink, and crucial insights on the dynamics of its growth have been obtained using capsule-binding proteins such as specific antibodies or complement. We have developed an alternative method that allows both static and time-lapse imaging of the capsule using Percoll^®, a suspension of nanometric spheres that do not penetrate the capsule. Given that these particles have a higher refractive index than the capsule, the latter can be imaged by differential interference contrast (DIC) microscopy. Static observation of the capsule with DIC and Percoll^® results in capsule thickness measurements that match those made with India ink. Using capsule-inducing media, a glass-bottom incubation chamber and a live-imaging system equipped for DIC microscopy, this method allows time-lapse imaging of capsule growth. In contrast with India ink staining, DIC imaging of Percoll^® exclusion halos result in crisp images. The greatest advantage of this method, though, is that unlike India ink, the Percoll^® particles are non-toxic and unlike opsonins they do not bind the capsule, resulting in observations of capsule growth that are free from interference of bound proteins on capsule physiology.

A protocol was developed to visualize and analyze the structure of membrane vesicles (MVs) from Gram-negative bacteria. It is now accepted that these micrometric spherical vesicles are commonly produced by cells from all three domains of life, so the protocol could be useful in the study of vesicles produced by eukaryotes and archaea as well as bacteria. The multiplicity of functions performed by MVs, related to cell communication, interaction with the immune system, pathogenesis, and nutrient acquisition, among others, has made MVs a hot topic of research.

Due to their small size (25-300 nm), the observation of MVs requires electron microscopy and is usually performed by transmission electron microscopy (TEM) of negatively stained MVs. Other protocols applied for their visualization include scanning electron microscopy, TEM after fixation and embedding of vesicles, or even atomic force microscopy. In some of these techniques, vesicle structure is altered by drying, while others are time-consuming and most of them can generate artifacts. Cryo-TEM after plunge freezing allows the visualization of samples embedded in a thin film of vitreous ice, which preserves their native cellular structures and provides the highest available resolution for the imaging. This is achieved by very high cooling rates that turn the intrinsic water of cells into vitreous ice, avoiding crystal formation and phase segregation between water and solutes. In addition to other types of characterization, an accurate knowledge of MV structure, which can be obtained by this protocol, is essential for MV application in different fields.

All bacteria, fungi and plant cells are surrounded by a cell wall. This complex network of polysaccharides and glycoproteins provides mechanical support, defines cell shape, controls cell growth and influences the exchange of substances between the cell and its surroundings. Despite its name, the cell wall is a flexible, dynamic structure. However, due to the lack of non-invasive methods to probe the structure, relatively little is known about the synthesis and dynamic remodeling of cell walls. Here, we describe a non-invasive method that quantifies a key physiological parameter of cell walls, the porosity, i.e., the size of spaces between cell wall components. This method measures the porosity-dependent decrease of the plasma membrane-localized fluorescent dye FM4-64 in the presence of the extracellular quencher Trypan blue. This method is applied to bacteria, fungi and plant cell walls to detect dynamic changes of porosity in response to environmental cues.

DNA damage repair proteins form foci in response to DNA damaging agents. The efficiency and integrity of the DNA repair pathway of a particular eukaryotic (mutant) strain is usually determined by the number of foci formed compared with their wild-type counterpart. Conventionally, focus number is determined visually, and this low accuracy may obscure the identification of a weaker phenotype, particularly when the output is low. Here, using the homologous recombination protein Rhp54 as an example, we present a protocol that can increase the consistency of foci identification among samples and can significantly improve the efficiency of foci quantification for large sample sizes. A similar method can be applied to other foci-forming proteins.

The evaluation of protein localization changes in cells under diverse chemical and genetic perturbations is now possible due to the increasing quantity of screens that systematically image thousands of proteins in an organism. Integrating information from different screens provides valuable contextual information about the protein function. For example, proteins that change localization in response to many different stressful environmental perturbations may have different roles than those that only change in response to a few. We developed, to our knowledge, the first protocol that permits the quantitative comparison and clustering of protein localization changes across multiple screens. Our analysis allows for the exploratory analysis of proteins according to their pattern of localization changes across many different perturbations, potentially discovering new roles by association.