微生物学

Microbial biofilms are structured communities of microorganisms embedded in a self-produced extracellular matrix, adhering to surfaces. These biofilms enhance bacterial resistance to antibiotics, immune responses, and environmental stress. Current microscopy techniques, such as scanning electron microscopy (SEM), confocal laser scanning microscopy (CLSM), and fluorescence microscopy, are commonly used to visualize and differentiate biofilms. However, their high cost and complexity often render them impractical. In contrast, simpler methods like crystal violet and Congo red staining are limited in distinguishing bacterial cells from the biofilm matrix. This study introduces a cost-effective, dual-staining method using Maneval’s stain to visualize and differentiate microbial biofilms. It requires only basic equipment and minimal reagents, making it ideal for routine use in clinical diagnosis and microbial research.

Capturing produced, consumed, or exchanged metabolites (metabolomics) and the result of gene expression (transcriptomics) require the extraction of metabolites and RNA. Multi-omics approaches and, notably, the combination of metabolomics and transcriptomic analyses are required for understanding the functional changes and adaptation of microorganisms to different physico-chemical and environmental conditions. A protocol was developed to extract total RNA and metabolites from less than 6 mg of a kind of phototrophic biofilm: oxygenic photogranules. These granules are aggregates of several hundred micrometers up to several millimeters. They harbor heterotrophic bacteria and phototrophs. After a common step for cell disruption by bead-beating, a part of the volume was recovered for RNA extraction, and the other half was used for the methanol- and dichloromethane-based extraction of metabolites. The solvents enabled the separation of two phases (aqueous and lipid) containing hydrophilic and lipophilic metabolites, respectively. The ¹H nuclear magnetic resonance (NMR) analysis of these extracts produced spectra that contained over a hundred signals with a signal-to-noise ratio higher than 10. The quality of the spectra enabled the identification of dozens of metabolites per sample. Total RNA was purified using a commercially available kit, yielding sufficient concentration and quality for metatranscriptomic analysis. This novel method enables the co-extraction of RNA and metabolites from the same sample, as opposed to the parallel extraction from two samples. Using the same sample for both extractions is particularly advantageous when working with inherently heterogeneous complex biofilm. In heterogeneous systems, differences between samples may be substantial. The co-extraction will enable a holistic analysis of the metabolomics and metatranscriptomics data generated, minimizing experimental biases, including technical variations and, notably, biological variability. As a result, it will ensure more robust multi-omics analyses, particularly by improving the correlation between metabolic changes and transcript modifications.

Campylobacter jejuni, a widespread pathogen found in birds and mammals, poses a significant risk for zoonosis worldwide despite its susceptibility to environmental and food-processing stressors. One of its main survival mechanisms is the formation of biofilms that can withstand various food-processing stressors, which is why efficient methods for assessing biofilms are crucial. Existing methods, including the classical culture-based plate counting method, biomass-staining methods (e.g., crystal violet and safranin), DNA-staining methods, those that use metabolic substrates to detect live bacteria (e.g., tetrazolium salts and resazurin), immunofluorescence with flow cytometry or fluorescence microscopy, and PCR-based methods for quantification of bacterial DNA, are diverse but often lack specificity, sensitivity, and suitability. In response to these limitations, we propose an innovative approach using NanoLuc as a reporter protein. The established protocol involves growing biofilms in microtiter plates, washing unattached cells, adding Nano-Glo luciferase substrate, and measuring bioluminescence. The bacterial concentrations in the biofilms are calculated by linear regression based on the calibration curve generated with known cell concentrations. The NanoLuc protein offers a number of advantages, such as its small size, temperature stability, and highly efficient bioluminescence, enabling rapid, non-invasive, and comprehensive assessment of biofilms together with quantification of a wide range of cell states. Although this method is limited to laboratory use due to the involvement of genetically modified organisms, it provides valuable insights into C. jejuni biofilm dynamics that could indirectly help in the development of improved food safety measures.

Candida auris, labeled an urgent threat by the CDC, shows significant resilience to treatments and disinfectants via biofilm formation, complicating treatment/disease management. The inconsistencies in biofilm architecture observed across studies hinder the understanding of its role in pathogenesis. Our novel in vitro technique cultivates C. auris biofilms on gelatin-coated coverslips, reliably producing multilayer biofilms with extracellular polymeric substances (EPS). This method, applicable to other Candida species like C. glabrata and C. albicans, is cost-effective and mimics the niche of biofilm formation. It is suitable for high-throughput drug screening and repurposing efforts, aiding in the development of new therapeutics. Our technique represents a significant advancement in Candida biofilm research, addressing the need for consistent, reproducible biofilm models. We detail a step-by-step procedure for creating a substratum for biofilm growth and measuring biofilm thickness using confocal laser scanning microscopy (CLSM) and ultrastructure by scanning electron microscopy (SEM). This method provides consistent outcomes across various Candida species.

The human pathogenic yeast Candida albicans can attach to epithelial cells or indwelling medical devices to form biofilms. These microbial communities are highly problematic in the clinic as they reduce both sensitivity to antifungal drugs and detection of fungi by the immune system. Amyloid structures are highly organized quaternary structures that play a critical role in biofilm establishment by allowing fungal cells to adhere to each other. Thus, fungal amyloids are exciting targets to develop new antifungal strategies. Thioflavin T is a specific fluorescent dye widely used to study amyloid properties of target proteins in vitro (spectrophotometry) and in vivo (epifluorescence/confocal microscopy). Notably, thioflavin T has been used to demonstrate the ability of Als5, a C. albicans adhesin, to form an amyloid fiber upon adhesion. We have developed a pipeline that allows us to study amyloid properties of target proteins using thioflavin T staining in vitro and in vivo, as well as in intact fungal biofilms. In brief, we used thioflavin T to sequentially stain (i) amyloid peptides, (ii) recombinant proteins, (iii) fungal cells treated or not with amyloid peptides, (iv) fungal amyloids enriched by cell fractionation, and (v) intact biofilms of C. albicans. Contrary to other methods, our pipeline gives a complete picture of the amyloid behavior of target proteins, from in vitro analysis to intact fungal biofilms. Using this pipeline will allow an assessment of the relevance of the in vitro results in cells and the impact of amyloids on the development and/or maintenance of fungal biofilm.

Key features

• Study of amyloid properties of fungal proteins.

• Visualization of the subcellular localization of fungal amyloid material using epifluorescence or confocal microscopy.

• Unraveling of the amyloid properties of target proteins and their physiological meaning for biofilm formation.

• Observation of the presence of amyloid structures with live-cell imaging on intact fungal biofilm using confocal microscopy.

Graphical overview

Campylobacter jejuni, a zoonotic foodborne pathogen, is the worldwide leading cause of acute human bacterial gastroenteritis. Biofilms are a significant reservoir for survival and transmission of this pathogen, contributing to its overall antimicrobial resistance. Natural compounds such as essential oils, phytochemicals, polyphenolic extracts, and D-amino acids have been shown to have the potential to control biofilms formed by bacteria, including Campylobacter spp. This work presents a proposed guideline for assessing and characterizing bacterial biofilm formation in the presence of naturally occurring inhibitory molecules using C. jejuni as a model. The following protocols describe: i) biofilm formation inhibition assay, designed to assess the ability of naturally occurring molecules to inhibit the formation of biofilms; ii) biofilm dispersal assay, to assess the ability of naturally occurring inhibitory molecules to eradicate established biofilms; iii) confocal laser scanning microscopy (CLSM), to evaluate bacterial viability in biofilms after treatment with naturally occurring inhibitory molecules and to study the structured appearance (or architecture) of biofilm before and after treatment.

Microbiologists are learning to appreciate the importance of “functional amyloids” that are produced by numerous bacterial species and have impacts beyond the microbial world. These structures are used by bacteria to link together, presumably to increase survival, protect against harsh conditions, and perhaps to influence cell-cell communication. Bacterial functional amyloids are also beginning to be appreciated in the context of host-pathogen interactions, where there is evidence that they can trigger the innate immune system and are recognized as non-self-molecular patterns. The characteristic three-dimensional fold of amyloids renders them similar across the bacterial kingdom and into the eukaryotic world, where amyloid proteins can be undesirable and have pathological consequences. The bacterial protein curli, produced by pathogenic Salmonella enterica and Escherichia coli strains, was one of the first functional amyloids discovered. Curli have since been well characterized in terms of function, and we are just starting to scratch the surface about their potential impact on eukaryotic hosts. In this manuscript, we present step-by-step protocols with pictures showing how to purify these bacterial surface structures. We have described the purification process from S. enterica, acknowledging that the same method can be applied to E. coli. In addition, we describe methods for detection of curli within animal tissues (i.e., GI tract) and discuss purifying curli intermediates in a S. enterica msbB mutant strain as they are more cytotoxic than mature curli fibrils. Some of these methods were first described elsewhere, but we wanted to assemble them together in more detail to make it easier for researchers who want to purify curli for use in biological experiments. Our aim is to provide methods that are useful for specialists and non-specialists as bacterial amyloids become of increasing importance.

Biofilms serve as a bacterial survival strategy, allowing bacteria to persist under adverse environmental conditions. The non-pathogenic Listeria innocua is used as a surrogate organism for the foodborne pathogen Listeria monocytogenes, because they share genetic and physiological similarities and can be used in a Biosafety Level 1 laboratory. Several methods are used to evaluate biofilms, including different approaches to determine biofilm biomass or culturability, viability, metabolic activity, or other microbial community properties. Routinely used methods for biofilm assay include the classical culture-based plate counting method, biomass staining methods (e.g., crystal violet and safranin red), DNA staining methods (e.g., Syto 9), methods that use metabolic substrates to detect live bacteria (e.g., tetrazolium salts or resazurin), and PCR-based methods to quantify bacterial DNA. The NanoLuc (Nluc) luciferase biofilm assay is a viable alternative or complement to existing methods. Functional Nluc was expressed in L. innocua using the nisin-inducible expression system and bacterial detection was performed using furimazine as substrate. Concentration dependent bioluminescence signals were obtained over a concentration range greater than three log units. The Nluc bioluminescence method allows absolute quantification of bacterial cells, has high sensitivity, broad range, good day-to-day repeatability, and good precision with acceptable accuracy. The advantages of Nluc bioluminescence also include direct detection, absolute cell quantification, and rapid execution.

Graphic abstract:

Engineering Listeria innocua to express NanoLuc and its application in bioluminescence assay.

Bacterial swarming refers to a rapid spread, with coordinated motion, of flagellated bacteria on a semi-solid surface (Harshey, 2003). There has been extensive study on this particular mode of motility because of its interesting biological and physical relevance, e.g., enhanced antibiotic resistance (Kearns, 2010) and turbulent collective motion (Steager et al., 2008). Commercial equipment for the live recording of swarm expansion can easily cost tens of thousands of dollars (Morales-Soto et al., 2015); yet, often the conditions are not accurately controlled, resulting in poor robustness and a lack of reproducibility. Here, we describe a reliable design and operations protocol to perform reproducible bacterial swarming assays using time-lapse photography. This protocol consists of three main steps: 1) building a “homemade,” environment-controlled photographing incubator; 2) performing a bacterial swarming assay; and 3) calculating the swarming rate from serial photos taken over time. An efficient way of calculating the bacterial swarming rate is crucial in performing swarming phenotype-related studies, e.g., screening swarming-deficient isogenic mutant strains. The incubator is economical, easy to operate, and has a wide range of applications. In fact, this system can be applied to many slowly evolving processes, such as biofilm formation and fungal growth, which need to be monitored by camera under a controlled temperature and ambient humidity.

Characterization of biofilm formation and metabolic activities is critical to investigating biofilm interactions with environmental factors and illustrating biofilm regulatory mechanisms. An appropriate in vitro model that mimics biofilm in vivo habitats therefore demands accurate quantitation and investigation of biofilm-associated activities. Current methodologies commonly involve static biofilm setups (such as biofilm assays in microplates, bead biofilms, or biofilms on glass-slides) and fluidic flow biofilm systems (such as drip-flow biofilm reactors, 3-channel biofilm reactors, or tubing biofilm reactors). Continuous flow systems take into consideration the contribution of hydrodynamic shear forces, nutrient supply, and physical transport of dispersed cells, which define the habitat for biofilm development in most natural and engineered systems. This protocol describes the assembly of 3 flow-system setups to cultivate Pseudomonas aeruginosa PAO1 and Shewanella oneidensis MR-1 model biofilms, including the respective quantitation and observation approaches. The standardized flow systems promise productive and reproducible biofilm experimental results, which can be further modified according to specific research projects.