微生物学

Drug biotransformation by the host microbiome can impact the therapeutic success of treatment. In the context of cancer, drug degradation can take place within the microenvironment of the targeted tumor by intratumor bacteria. In pancreatic cancer, increased chemo-resistance against the frontline chemotherapy gemcitabine is thought to arise from drug degradation by the tumor microbiome. This bacterial–drug interaction highlights the need for developing rapid assays for monitoring bacterial gemcitabine breakdown. While chemical approaches such as high-performance liquid chromatography are suitable for this task, they require specialized equipment and expertise and are limited in throughput. Functional cell-based assays represent an alternate approach for performing this task. We developed a functional assay to monitor the rate of bacterial gemcitabine breakdown using a highly sensitive bacterial reporter strain. Our method relies on standard laboratory equipment and can be implemented at high throughput to monitor drug breakdown by hundreds of strains simultaneously. This functional assay can be readily adapted to monitor degradation of other drugs.

Key features

• Quantification of gemcitabine breakdown by incubating bacteria that degrades the drug and subsequently testing the growth of a reporter strain on filtered supernatant.

• Use of an optimized reporter strain that was genetically engineered to be a non-degrader strain and highly sensitive to gemcitabine.

• A high-throughput assay performed in microplates that can be adjusted for identifying bacteria with a fast or slow gemcitabine degradation rate.

• The assay results can be compared to results from a standard curve with known drug concentrations to quantify degradation rate.

Graphical overview

Protocol overview. (1) Bacteria are incubated with gemcitabine for a set period of time. (2) Samples are removed from co-incubated suspensions and filtered to remove bacteria to halt gemcitabine degradation. (3) A gemcitabine-sensitive reporter strain is then added to the conditioned supernatant and is supplemented with growth media. (4) Growth of the reporter strain is monitored over time. (5) Results from the growth experiments are used to infer the concentration of gemcitabine in the co-culture supernatant and the drug degradation rate.

Bacterial studies based on growth curves are common in microbiology and related fields. Compared to the standard photometer and cuvette based protocols, bacterial growth curve measurements with microplate readers provide better temporal resolution, higher efficiency, and are less laborious, while analysis and interpretation of the microplate-based measurements are less straightforward. Recently, we developed a new analysis method for evaluating bacterial growth with microplate readers based on time derivatives. Here, we describe a detailed protocol for this development and provide the homemade program for the new analysis method.

Control of malaria caused by Plasmodium vivax can be improved by the discovery and development of novel drugs against the parasite’s liver stage, which includes relapse-causing hypnozoites. Several recent reports describe breakthroughs in the culture of the P. vivax liver stage in 384-well microtiter plates, with the goal of enabling a hypnozoite-focused drug screen. Herein we describe assay details, protocol developments, and different assay formats to interrogate the chemical sensitivity of the P. vivax liver stage in one such medium-throughput platform. The general assay protocol includes seeding of primary human hepatocytes which are infected with P. vivax sporozoites generated from the feeding of Anopheles dirus mosquitoes on patient isolate bloodmeals. This protocol is unique in that, after source drug plates are supplied, all culture-work steps have been optimized to preclude the need for automated liquid handling, thereby allowing the assay to be performed within resource-limited laboratories in malaria-endemic countries. Throughput is enhanced as complex culture methods, such as extracellular matrix overlays, multiple cell types in co-culture, or hepatic spheroids, are excluded as the workflow consists entirely of routine culture methods for adherent cells. Furthermore, installation of a high-content imager at the study site enables assay data to be read and transmitted with minimal logistical delays. Herein we detail distinct assay improvements which increase data quality, provide a means to limit the confounding effect of hepatic metabolism on assay data, and detect activity of compounds with a slow-clearance phenotype.

Graphical abstract:

Overview of P. vivax liver stage screening assay performed at the Institute Pasteur of Cambodia.

Antimicrobial-resistant Mycobacterium tuberculosis (Mtb) causes over 200,000 deaths globally each year. Current assays of antimicrobial resistance require knowledge of the mutations that confer drug resistance or long periods of culture time to test growth under drug pressure. We present ODELAM (One-cell Doubling Evaluation of Living Arrays of Mycobacterium), a time-lapse microscopy-based method that observes individual cells growing into microcolonies. This protocol describes sample and media preparation and contains instructions for assembling the ODELAM sample chamber. The ODELAM sample chamber is designed to provide a controlled environment to safely observe the growth of Mtb by time-lapse microscopy on an inverted wide-field microscope. A brief description of the ODELAM software is also provided here. ODELAM tracks up to 1500 colony forming units per region of interest and can observe up to 96 regions for up to seven days in a single experiment. This technique allows the quantification of population heterogeneity. ODELAM enables rapid quantitative measurements of growth kinetics in as few as 30 h under a wide variety of environmental conditions.

Graphic abstract:

Schematic representation of the ODELAM platform

Lysogenic phages can integrate into their bacterial host’s genome, potentially transferring any genetic information they possess including virulence or resistance genes, and are therefore routinely excluded from therapeutic applications. Lysogenic behavior is typically seen in phages that create turbid plaques or possess subpar bactericidal activity; yet, these are not definitive indicators. As a result, the presence of integrase genes is often used as a hallmark for lysogenic behavior; however, the accuracy of genetic screening for lysogeny depends on the quality of the extraction, sequencing and assembly of the phage genome, and database comparison. The present protocol describes a simple phenotypic test that can be used to screen therapeutically relevant phages for lysogenic behavior. This test relies on the identification of spontaneous phage release from their lysogenized host and can be reliably used in cases where no sequencing data are available. The protocol does not require specialized equipment, is not work-intensive, and is broadly applicable to any phage with an easily culturable bacterial host, making it particularly amenable to settings with limited resources.

Graphical abstract:

Screening pipeline for lysogen activity of a given phage

Antibacterial coatings have currently gained great importance in biomedical technology investigations. Because of the spatial arrangement of the film coatings, evaluation of antibacterial activity presents a new challenge regarding traditional bacterial counting methods. In this protocol, four clinically relevant pathogens, Salmonella typhimurium, Escherichia coli, Pseudomonas aeruginosa, and Staphylococcus aureus were incubated on titania mesostructured thin film coatings for 24 h. Then, cell viability was studied considering three methods: counting of the number of colony forming units (CFU), live/dead staining, and quantification of extracellular DNA in suspension. Firstly, bacterial count was determined by the standard plate-count technique. Secondly, bacteria membrane integrity was evaluated by utilization of two fluorescent dyes, which allow distinction between live (membrane intact) and dead (membrane disrupted) bacteria. Lastly, extracellular DNA was quantified by spectrophotometry. In this manner, the three aforementioned techniques enabled the study of bacterial viability by qualitative and quantitative analyses.

Bdellovibrio bacteriovorus, an obligate predatory bacterium [i.e., bacteria that kill and feed on other bacteria (prey)], has the potential to be used as a probiotic for the disinfection of surfaces or for the treatment of bacterial infections. One option is to use this organism in combination with antimicrobials to potentiate the effectiveness of treatments. In order to make this approach feasible more has to be known about the ability of B. bacteriovorus to resist antibiotics itself. Standard assays to determine the minimum inhibitory concentration (MIC) are not suitable for B. bacteriovorus, since the small size of this bacterium (0.25-0.35 by 0.5-2 μm) prevents scattering at OD₆₀₀. Since these predatory bacteria require larger prey bacteria for growth (e.g., E. coli dimensions are 1 by 1-2 μm), the basis for the antimicrobial sensitivity assay described here is the reduction of the OD₆₀₀ caused by prey lysis during growth. Previous studies on predatory bacteria resistance to antimicrobials employed methods that did not allow a direct comparison of antimicrobial resistance levels to those of other bacterial species. Here, we describe a procedure to determine B. bacteriovorus sensitivity to antimicrobials which can be compared to a reference organism tested as close as possible to the same experimental conditions. Briefly, minimal inhibitory concentration (MIC) values of B. bacteriovorus are determined by measuring the reduction in absorbance at 600 nm of mixed predator/prey cultures in presence and absence of different antimicrobial concentrations. Of note, this method can be modified to obtain antimicrobial MIC values of other predatory bacteria, using different conditions, prey bacteria and/or antimicrobials.

Bacteriocins are small ribosomally synthesized antimicrobial peptides produced by some microorganisms including lactic acid bacteria (LAB), a group of Gram-positive bacteria (cocci, rods) expressing high tolerance for low pH. Bacteriocins kill bacteria rapidly and are biologically active at very low concentrations. Bacteriocins produced by LAB are primarily active against closely related bacterial species. Many bacteriocins have been investigated with respect to their potential use in promoting human, plant, and animal health, and as food biopreservatives. Bacteriocins produced by LAB are particularly interesting since several LAB have been granted GRAS (Generally Recognized as Safe) status. Because it is not always possible to extract active bacteriocins secreted from cells grown in liquid medium, we developed a simple and inexpensive peptide extraction procedure using a semi-solid nutrient-rich agar medium. We hereby present a detailed procedure that leads to the rapid extraction of secreted bioactive bacteriocin peptides from the oral species Streptococcus mutans, a prolific bacteriocin-producing species, and its potential application for bacteriocin extraction from other LAB (e.g., Streptococcus, Lactococcus, Enterococcus). We also present a simple method for the detection of bacteriocin activity from the purified extracellular peptide extract.

This protocol details the construction of a simple, low-cost, small-scale, multiplex chemostat system designed for the continuous cultivation of microorganisms in suspension (i.e., bacteria, yeast, microalgae). The continuous culture device can operate at a working volume of 25 ml and allows the run of 8 chemostats in parallel by a single person. It provides a platform for parallel, long-term studies of evolution and adaptation of microorganisms under the stress of antimicrobial agents and/or toxic pollutants. The system complies with the varied needs of researchers for an accessible, highly-throughput and reliable tool that is nevertheless easy to construct, use and operate, and not demanding of space, materials, medium supply and workload. Here, we also validate the use of this system to generate de novo resistance towards a novel antimicrobial and a commonly used antibiotic in an antimicrobial-sensitive model organism. We believe that this "Do It Yourself" (DIY) system may constitute a useful tool to address the global problem of antibiotic resistance and to develop non-antibiotic based therapies.

The emergence and rapid spread of multidrug resistance in bacteria have led to the urgent need for novel antibacterial agents. Membrane permeabilization is the mechanism for many antibacterial molecules that are being developed against gram-negative bacteria. Thus, to determine the efficacy of a potential antibacterial molecule, it is important to assess the change in bacterial membrane permeability after treatment. This study describes the protocol for the assays of outer and inner membrane permeability using the fluorescent probes N-phenyl-1-naphthylamine and propidium iodide. Compared with other experiments, such as electron microscopy and the assay of minimal bactericidal concentration, this methodology provides a simpler, faster, and cost-effective way of estimating the membrane-permeabilizing effect and bactericidal efficacy of antibacterial molecules. This study presents an optimized protocol with respect to the classical protocols by incubating bacteria with antibacterial molecules in the culture condition identical to that of antibacterial assays and then detecting the signal of the fluorescent probe in the buffer without broth and antibacterial molecules. This protocol avoids the effect of nutrient deficiency on the physiological status of bacteria and the interference of antibacterial molecules towards the fluorescent probe. Thus, this method can effectively and precisely evaluate the membrane permeability and match the results obtained from other antibacterial assays, such as minimum inhibitory concentration and time–kill curve assays.