Microbiology

Inteins are elements translated within host proteins and removed via a unique protein splicing reaction. In this process, the two peptide bonds flanking the intein are rearranged, releasing the intein and leaving a standard peptide bond in its place. Due to their ability to shuffle peptide bonds in a specific and controlled manner, inteins have proven valuable in protein engineering, leading to the development of numerous impactful technologies. In one application, intein-based biosensors link the activity of a host protein to intein excision. Recently, we developed a biosensor to measure protein stability in vivo, in which the removal of an intein-protein fusion is required for antibiotic resistance. In our protocol, cells expressing our biosensor are logarithmically diluted and spotted on agar plates containing increasing levels of antibiotics. Following incubation, quantitative survival curves can be generated. We also developed a dual protein stability sensor where both antibiotic resistance and fluorescence can be used as readouts and demonstrated that co-expression of the chaperonin GroEL can promote survival and fluorescence. Taken together, our novel intein-based biosensor adds to the available tools to measure protein stability within the cellular environment.

Reverse genetics systems in virology are technologies used to generate recombinant viruses, enabling the manipulation of viral genes. Recombinant viruses facilitate the investigation of pathogenesis and the development of antivirals. In studies of positive-sense single-stranded RNA (ssRNA) viruses, a reverse genetics approach typically uses infectious viral cDNA clones derived from bacterial artificial chromosomes and plasmids or from the in vitro ligation of viral cDNA fragments. However, these methods are time-consuming, involve complex procedures, and do not always successfully generate recombinant viruses. Possible reasons for unsuccessful outcomes include i) viral sequences exhibiting toxicity in bacterial systems, ii) the duplication of viral genes observed in some strains, complicating the acquisition of correct cDNA clones, and iii) certain cell lines being highly susceptible to infection but difficult to transfect with nucleotides. For these reasons, a simple and rapid reverse genetics system is needed to accelerate research on ssRNA viruses. The circular polymerase extension reaction (CPER) method offers a solution by eliminating the need for molecular cloning in bacteria, enabling the generation of recombinant viruses over a shorter timeframe. This method has been widely adopted for the study of ssRNA viruses, including SARS-CoV-2 and flaviviruses. Recently, we expanded the CPER method for ssRNA viruses using internal ribosome entry site (IRES)-mediated translation. This protocol details the experimental procedures, using bovine viral diarrhea virus as an example—one of the most challenging viruses for generating viral cDNA clones because of the factors listed above.

Quiescence, the temporary and reversible exit from proliferative growth, is a fundamental biological process. Budding yeast is a preeminent model for studying cellular quiescence owing to its rich experimental toolboxes and evolutionary conservation across eukaryotic pathways and processes that control quiescence. Yeast quiescent cells are reported to be isolated by the continuous linear Percoll gradient method and identified by combining different features such as cell cycle, heat resistance, and cell morphology (single cell). Generally, 10–25 mL of Percoll isotonic solution is first obtained by mixing Percoll with NaCl in 12.5–30 mL centrifugal tubes. Then, the gradient is prepared at high speed for 15–60 min. Finally, approximately 2 × 10⁹ cells are collected, overlaid onto the preformed gradient, and centrifuged to obtain distinct cell fractions. This method requires more reagents and samples and special centrifuges and centrifuge tubes. Besides the cost, it is less favorable for experiments that require high-throughput analyses with a small volume of sample each time. The protocol described here aims to solve those problems by combining the use of 2 mL centrifugal tubes with density marker beads. The protocol also focuses on how to optimize the buoyant density distribution of the density gradient solution such that the density bands better match those of different fraction cells. This will help fully separate quiescent and non-quiescent cells. The protocol can be easily adapted to a wide variety of unicellular microbes with different buoyancy density differentiation during cultivation, such as yeast and bacteria.

Xylan is the main component of hemicellulose and consists of a complex heteropolysaccharide with a heterogeneous structure. This framework, in addition to the crystalline structure of cellulosic fibers and the rigidity of lignin, makes lignocellulosic biomass (LCB) highly recalcitrant to degradation. Xylanases are glycoside hydrolases that cleave the β-1,4-glycoside linkages in the xylan backbone and have attracted increasing attention due to their potential uses in various industrial sectors such as pulp and paper, baking, pharmaceuticals, and lignocellulosic biorefining. For decades, the measurement of xylanase activity was based on reducing sugar quantification methods like DNS or Nelson/Somogyi assays, with numerous limitations in terms of specificity and interference from other enzymatic activities. A better alternative is the colorimetric Azo-Xylan assay, which specifically measures the endo-1,4-β-D-xylanase activity. In this study, the Azo-Xylan protocol was adapted from the company Megazyme to determine the enzymatic activity of thermostable xylanases produced by microbial consortia (i.e., microbiomes), aiming to determine biochemical features such as temperature and pH optima, thermostability, and shelf life. This modified approach offers a rapid, cost-effective, and highly specific method for the determination of xylanase activity in complex mixtures, helping the development of a xylanase-based method for the hydrolysis of hard-degrading substrates in bio-based industries.

The ability to efficiently screen plant pathogen effectors is crucial for understanding plant–pathogen interactions and developing disease-resistant crops. Traditional methods are often labor-intensive and time-consuming. Here, we present a robust, high-throughput screening assay using the tobacco mosaic virus–green fluorescent protein (TMV-GFP) vector system. The screening system combines the TMV-GFP vector and Agrobacterium-mediated transient expression in the model plant Nicotiana benthamiana. This system enables the rapid identification of effectors that interfere with plant immunity (both activation and suppression). The biological function of these effectors can be easily evaluated within six days by observing the GFP fluorescence signal using a UV lamp. This protocol significantly reduces the time required for screening and increases the throughput, making it suitable for large-scale studies. The method is versatile, cost-effective, and can be adapted to effectors with immune interference activity from various pathogens.

In molecular diagnosis, DNA extraction kits are sample-specific and proprietary, preventing lateral distribution among similar facilities from different sectors to alleviate supply shortages during a crisis. Previous fast extraction protocols such as detergent-based ones allow fast DNA extraction for nucleic acid amplification tests (NAAT), mainly polymerase chain reaction (PCR). The use of NaOH (dense alkali) to rupture cells and nuclei and destabilize the conformation of DNases might alleviate shortages and costs while retaining enough robustness to treat complicated samples with minimal environmental and logistical footprint. Biological samples are hand-crushed using a pestle in 1.5 mL tubes with 360 μL of 0.2 M NaOH for 3–5 min and incubated at 75 °C for 10 min. For immediate use, 115.2 μL of 1 M Tris (pH 8) and 364.8 μL nuclease-free water are added, and the sample is vortexed for 10 s and spun at 10,000× g for 3 min; then, 700 μL is transferred to a clean microtube. Two serial dilutions follow, and all concentrations are used as templates for PCR. A refined, storable extract can be produced by adding 70 μL of HCl 1 M (instead of Tris-HCl) and one volume of cold isopropanol to the extract for standard precipitation. This method can increase throughput in emergencies by field deployment in resource-limited settings (RLS) or allow benchtop backup in cases of acquisition disruption or sample surge in established facilities. The crude extract can be used for immediate PCR in both benchtop and portable thermocyclers, thus allowing NAAT in resource-limited settings with low costs and waste footprint or during prolonged crises, where supply chain failures may occur. The refined version produces alcohol-precipitated nucleic acids, suitable for both immediate use and for storage or dispatch for spatiotemporally separate analysis while offering much better amplification quality with a small increase in time and minimal increase in expendables/chemicals needed.

Laboratory-developed tests (LDTs) are optimal molecular diagnostic modalities in circumstances such as public health emergencies, rare disease diagnosis, limited budget, or where existing commercial alternatives are unavailable, limited in supply, or withdrawn, either temporarily or permanently. These tests reduce access barriers and enhance equitable clinical practice and healthcare delivery. Despite recommendations for the development of nucleic acid amplification tests, procedural details are often insufficient, inconsistent, and arbitrary. This protocol elucidates the methodology used in the development of a fully automated real-time polymerase chain reaction (qPCR)-based test, using the Panther Fusion^® Open Access^TM functionality, for the detection of Streptococcus agalactiae in pregnant women, using selectively enriched rectovaginal swabs. In addition, guidelines are provided for oligonucleotide design (primers and TaqMan probes), in silico and in vitro evaluation of design effectiveness, optimization of the physicochemical conditions of the amplification reaction, and result analysis based on experimental designs and acceptance criteria. Furthermore, recommendations are provided for the analytical and clinical validation of the intended use. Our approach is cost-effective, particularly during the design and optimization phases. We primarily used open-source bioinformatics software and tools for in silico evaluations for the test design. Subsequently, the process was manually optimized using a CFX96 Dx analyzer, whose technical specifications and performance are homologous to that of the final platform (Panther Fusion^®). Unlike Panther Fusion^®, the CFX96 Dx does not require excess volumes of reagents, samples, and evaluation materials (dead volume) to accommodate potential robotic handling-associated imprecisions. The utilization of the CFX96 Dx analyzer represents a strategic approach to enhancing the efficiency of resources and the optimization of time during LDT optimization.

Enteroviruses are abundant pathogens of humans and animals. Their replication is strictly dependent on the conserved, viral AAA+ ATPase 2C. 2C is an oligomerizing, peripheral membrane protein, and its low solubility as recombinant protein has hampered functional studies of the full-length, recombinant protein bound to a membrane. Here, we describe a modification of the classical, ultracentrifugation-based liposome flotation assay optimized to study the interaction of recombinant 2C with membranes and the functions of membrane-bound, full-length recombinant 2C. The assay takes advantage of the high solubility of recombinant 2C while fused to a maltose-binding protein. Removing this solubility-enhancing tag by specific protease cleavage in the presence of liposomes allows 2C to associate with membranes prior to aggregating. Fluorophore labeling of protein and liposomes allows rapid and precise quantitation of 2C’s association with membranes. This assay is adaptable to any peripheral membrane protein that can be fluorophore-labeled and expressed as a solubility-enhancing fusion protein.

Protein palmitoylation is a lipid modification where a palmitoyl group is covalently attached via a thioester linkage to one or more cysteines on a substrate protein. This modification, catalyzed by a group of enzymes named DHHC enzymes after their conserved Asp-His-His-Cys motif, plays a significant role in regulating the localization, stability, and function of a wide range of cellular and viral proteins. By influencing how and where proteins interact within the cell, palmitoylation is essential for various cellular processes, including signaling pathways, membrane dynamics, and protein–protein interactions. Here, we describe the acyl-RAC assay, a biochemical technique designed to specifically enrich and analyze palmitoylated proteins from complex biological samples, such as cell lysates or tissue extracts. The assay begins by reducing and blocking free cysteine thiol groups on proteins, ensuring that only those thiols involved in thioester bonds with palmitates are accessible for downstream analysis. These thioester bonds are then cleaved to release the fatty acids from the cysteines, which are subsequently captured using thiopropyl Sepharose beads that bind to the newly exposed thiol groups. The captured proteins are eluted from the beads by breaking the bond between the thiol and the resin with reducing agents, and the proteins are then analyzed by SDS-PAGE followed by western blotting to identify and quantify them. The acyl-RAC assay's specificity for S-palmitoylated proteins makes it an invaluable tool for exploring this modification. It not only allows for the identification of previously unknown palmitoylated proteins, thereby deepening our understanding of palmitoylation in cellular processes and viral infections, but it also enables quantitative comparisons of protein palmitoylation under different experimental conditions or treatments.

The early detection of meningitis pathogens—including Haemophilus influenzae, Neisseria meningitidis, Streptococcus pneumoniae, and Klebsiella pneumoniae—through point-of-care (POC) systems is essential for mitigating the risk of neurological damage, enhancing patient outcomes, and facilitating prompt clinical decision-making. Nucleic acid amplification testing (NAAT) is a promising tool for improving the diagnosis process of bacterial pathogens associated with brain inflammation. This is due to its high sensitivity, rapidity, and compatibility with portable diagnostic platforms, making it particularly suitable for POC applications. This protocol introduces an innovative diagnostic approach designed to function effectively without the need for advanced laboratory equipment. By leveraging dual-priming isothermal amplification (DAMP), the assay uses custom internal primers to enhance specificity and minimize false results. Brilliant Green is used in this assay for fluorescence detection due to its availability, high fluorescence level, and optimal sample-to-background (S/B) ratio. The assay demonstrated excellent specificity, absence of false positives, sensitivity comparable to loop-mediated isothermal amplification (LAMP), and a high S/B ratio.