Biological Sciences


Protocols in Current Issue
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0 Q&A 278 Views Oct 5, 2023

Mixed communities of fungi and bacteria have been shown to be more efficient in degrading wood than fungi alone. Some standardised protocols for quantification of the wood decay ability of fungi have been developed (e.g., DIN V ENV 12038:2002 as the legal standard to test for the resistance of wood against wood-destroying basidiomycetes in Germany). Here, we describe a step-by-step protocol developed from the official standard DIN V ENV12038 to test combinations of bacteria and fungi for their combined wood degradation ability. Equally sized wood blocks are inoculated with wood decay fungi and bacterial strains. Axenic controls allow the analysis of varying degradation rates via comparison of the wood dry weights at the end of the experiments. This protocol provides new opportunities in exploration of inter- and intra-kingdom interactions in the wood-related environment and forms the basis for microcosm experiments.

Key features

• Quantification of wood decay ability of mixed cultures.

• Allows testing if fungi are more efficient in degrading wood when bacteria are present.

0 Q&A 388 Views Mar 20, 2023

Over the past decades, the main techniques used to visualize bacteria in tissue have improved but are still mainly based on indirect recognition of bacteria. Both microscopy and molecular recognition are being improved, but most procedures for bacteria detection in tissue involve extensive damage. Here, we describe a method to visualize bacteria in tissue slices from an in vivo model of breast cancer. This method allows examining trafficking and colonization of fluorescein-5-isothiocyanate (FITC)-stained bacteria in various tissues. The protocol provides direct visualization of fusobacterial colonization in breast cancer tissue. Rather than processing the tissue or confirming bacterial colonization by PCR or culture, the tissue is directly imaged using multiphoton microscopy. This direct visualization protocol causes no damage to the tissue; therefore, all structures can be identified. This method can be combined with others to co-visualize bacteria, types of cells, or protein expression in cells.

0 Q&A 2314 Views Jan 20, 2022

Caenorhabditis elegans is a ubiquitous free-living nematode that feeds on bacteria. The organism was introduced into a laboratory setting in the 1970s and has since gained popularity as a model to study host-bacteria interactions. One advantage of using C. elegans is that its intestine can be colonized by the bacteria on which it feeds. Quantifying the bacterial load within C. elegans is an important and easily obtainable metric when investigating host-bacteria interactions. Although quantification of bacteria harbored in C. elegans via whole-worm lysis is not a novel assay, there is great variation between existing methods. To lyse C. elegans, many protocols rely on the use of a hand-held homogenizer, which could introduce systematic error and subsequent variation between researchers performing the same experiment. Here, we describe a method of lysing the intestines of C. elegans to quantify the bacterial load within the intestine. Our method has been optimized for removing exogenous bacteria while maintaining worm paralysis, to ensure no bactericidal agents are swallowed, which could kill bacteria within the intestine and affect results. We utilize and compare the efficiency of two different homogenization tools: a battery-powered hand-held homogenizer, and a benchtop electric homogenizer, where the latter minimizes variability. Thus, our protocol has been optimized to reduce systematic error and decrease the potential for variability among experimenters.

Graphic abstract:

Simplified overview of the procedure used to quantify the bacterial load within C. elegans. The two different methods are herein described for worm lysis: “Option 1” is a hand-held homogenizer, and “Option 2” is a benchtop homogenizer.

0 Q&A 2073 Views Sep 20, 2021

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.

0 Q&A 1896 Views Aug 20, 2021

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.

0 Q&A 3946 Views May 5, 2021

Siderophores, a key substance that microorganisms produce to obtain iron under iron-limited conditions, play an important role in regulating interactions between beneficial bacteria and pathogenic bacteria. A large number of bacteria were isolated from the rhizosphere, and we used the method presented here to assay the siderophore production by these rhizosphere bacteria. This method is a modified version of the universal chrome azurol S (CAS) assay that uses a 96-channel manual pipetting workstation. By combining the liquid CAS assay with the multi-channel pipette workstation, high-throughput and rapid detection of siderophore production can be achieved. In summary, this method can be used to gain a general understanding of siderophore production by rhizosphere bacteria.

0 Q&A 2337 Views Oct 5, 2020
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.

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