Microbiology

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.

Inkjet 3D printing is an additive manufacturing method that allows the user to produce a small batch of customized devices for comparative study versus commercial products. Here, we describe the use of a commercial 2D ink development system (Dimatix material printing) to manufacture small batches of 3D medical or other devices using a recently characterized fungal anti-attachment material. Such printed devices may resist problems that beset commercial medical products due to colonization by the fungal pathogen Candida albicans. By sequentially introducing the cross-section bitmaps of the product’s CAD model and elevating the print head height using the auto-clicking script, we were able to create complex self-support geometries with the 2D ink development system. The use of this protocol allows researchers to produce a small batch of specimens for characterization from only a few grams of raw material. Additionally, we describe the testing of manufactured specimens for fungal anti-attachment. In comparison with most commercial AM systems, which require at least a few hundred grams of ink for printing trials, our protocol is well suited for smaller-scale production in material studies.

Development of biofilm associated candidemia for patients with implanted biomaterials causes an urgency to develop antimicrobial and biofilm inhibitive coatings in the management of recalcitrant Candida infections. Recently, there is an increase in the number of patients with biofilm formation and resistance to antifungal therapy. Therefore, there is a growing interest to use essential oils as coating agents in order to prevent biomaterial-associated Candida infections. Often high costs, complicated and laborious technologies are used for both applying the coating and determination of the antibiofilm effects hampering a rapid screening of essential oils. In order to determine biofilm formation of Candida on essential oil coated surfaces easier, cheaper and faster, we developed an essential oil (lemongrass oil) coated surface (silicone-rubber) by using a hypromellose ointment/essential oil mixture. Furthermore, we modified the “crystal violet binding assay” to quantify the biofilm mass of Candida biofilm formed on the lemongrass oil coated silicone rubber surface. The essential oil coating and the biomass determination of biofilms on silicone rubber can be easily applied with simple and accessible equipment, and will therefore provide rapid information about whether or not a particular essential oil is antiseptic, also when it is used as a coating agent.

Cyclic diguanylate monophosphate (c-di-GMP) is a second messenger signaling molecule that drives the transition from planktonic to the biofilm mode of growth in many bacterial species. Pseudomonas aeruginosa has at least two surface sensing systems that produce c-di-GMP in response to surface attachment, the Wsp and Pil-Chp systems. We recently used a plasmid-based c-di-GMP reporter (pP_cdrA::gfp) to describe how the Wsp system generates heterogeneity in surface sensing, resulting in two physiologically distinct subpopulations of cells during early biofilm formation. One subpopulation has elevated c-di-GMP and produces biofilm matrix, serving as the founders of initial microcolonies. The other subpopulation has low c-di-GMP and engages in surface motility, allowing for exploration of the surface. Here, we describe the protocol for a key experiment to confirm our initial observation of c-di-GMP heterogeneity during surface sensing: the use of flow-assisted cell sorting (FACS) to isolate subpopulations of cells with high and low c-di-GMP reporter activity, followed by quantitative Reverse Transcriptase PCR (qRT-PCR) of genes that are known to be transcriptionally regulated in response to cellular c-di-GMP levels (pelA, pslA). This protocol can be adapted by others to isolate subpopulations of high- and low- c-di-GMP P. aeruginosa cells that are genetically identical, but phenotypically distinct for future experiments examining specific mRNA transcripts as we did or, presumably, for additional applications like RNAseq, proteomics, or TNseq.

Graphical abstract

Bacterial surface adhesion, the first step in many important processes including biofilm formation and tissue invasion, is a fast process that occurs on a time scale of seconds. Adhesion patterns tend to be stochastic and spatially heterogeneous, especially when bacteria are present in low population densities and at early stages of adhesion to the surface. Thus, in order to observe this process, a high degree of temporal resolution is needed across a large surface area in a way that allows several replicates to be monitored. Some of the current methods used to measure bacterial adhesion include microscopy, staining-based microtiter assays, spectroscopy, and PCR. Each of these methods has advantages in assaying aspects of bacterial surface adhesion, but none can capture all features of the process. In the protocol presented here, adapted from Shteindel et al., 2019, fluorescently-labeled bacteria are monitored in a multi-titer setting using a standard plate fluorimeter and a dye that absorbs light in the fluorophore excitation and emission wavelengths. The advantage of using this dye is that it restricts the depth of the optic layer to the few microns adjacent to the bottom of the microtiter well, eliminating fluorescence originating from unattached bacteria. Another advantage of this method is that this setting does not require any preparatory steps, which enables reading of the sample to be repeated or continuous. The use of a standard multi-titer well allows easy manipulation and provides flexibility in experimental design.

Oxygenic photogranules (OPGs) are dense, three-dimensional aggregates containing a syntrophic, light-driven microbial community. Their temporal and spatial development interests microbial ecologists working at the bioprocess engineering interface, as this knowledge can be used to optimize biotechnological applications, such as wastewater treatment and biomass valorization. The method presented here enables the high-throughput quantification of photogranulation. OPGs are produced from a loose sludge-like microbial matrix in hydrostatic batch cultures exposed to light. This matrix transforms into a consolidated, roughly spherical aggregate over time. Photogranulation is quantified by time-lapse imaging coupled to automated image analysis. This allows studying the development of many OPGs simultaneously and in a fully automated way to systematically test what factors drive photogranulation. The protocol can also be used to quantify other types of (a)biotic aggregation.

Pseudomonas aeruginosa is a human pathogen capable to form robust biofilms. P. aeruginosa biofilms represent a serious problem because of the adverse effects on human health and industry, from sanitary and economic points of view. Typical strategies to break down biofilms have been long used, such as the use of disinfectants or antibiotics, but also, according to their high resistance to standard antimicrobial approaches, alternative strategies employing photocatalysis or control of biofilm formation by modifying surfaces, have been proposed. Colony forming units (cfu) counting and live/dead staining, two classic techniques used for biofilm quantification, are detailed in this work. Both methods assess cell viability, a key factor to analyze the microbial susceptibility to given treatment, then, they represent a good approach for evaluation of an antibiofilm strategy.