Journal of biological methods最新文献

DNA adenine methylation identification (DamID) is a powerful method to determine DNA binding profiles of proteins at a genomic scale. The method leverages the fusion between a protein of interest and the Dam methyltransferase of E. coli, which methylates proximal DNA in vivo. Here, we present an optimized procedure, which was developed for tissue-specific analyses in Caenorhabditis elegans and successfully used to footprint genes actively transcribed by RNA polymerases and to map transcription factor binding in gene regulatory regions. The present protocol details C. elegans-specific steps involved in the preparation of transgenic lines and genomic DNA samples, as well as broadly applicable steps for the DamID procedure, including the isolation of methylated DNA fragments, the preparation of multiplexed libraries, Nanopore sequencing, and data analysis. Two distinctive features of the approach are (i) the use of an efficient recombination-based strategy to selectively analyze rare cell types and (ii) the use of Nanopore sequencing, which streamlines the process. The method allows researchers to go from genomic DNA samples to sequencing results in less than a week, while being sensitive enough to report reliable DNA footprints in cell types as rare as 2 cells per animal.

Genomic manipulation offers the possibility for novel therapies in lieu of medical interventions in use today. The ability to genetically restore missing inflammatory genes will have a monumental impact on our current immunotherapy treatments. This study compared the efficacy of two different genetic manipulation techniques: clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR associated protein 9 (Cas9) transfection to adenoviral transduction to determine which method would provide the most transient and stable knockdown of myeloid differentiation primary response 88 (MyD88). MyD88 is a major regulator of nuclear factor kappa light chain enhancer of activated B cells (NFκB) pathway in Raw 264.7 macrophages. Following genetic manipulation, cells were treated for 24 h with Lipopolysaccharide (LPS) to stimulate the inflammatory pathway. Confirmation of knockdown was determined by western immunoblotting and quantification of band density. Both CRISPR/Cas9 and adenoviral transduction produced similar knockdown efficiency (~64% and 60%, respectively) in MyD88 protein 48 h post adenoviral transduction. NFκB phosphorylation was increased in CRISPR/Cas9-mediated MyD88 knockdown and control cells, but not in adenovirus-mediated MyD88 knockdown cells, following LPS administration. CRISPR/Cas9-mediated MyD88 knockdown macrophages treated with LPS for 24 h showed a 65% reduction in tumor necrosis factor alpha (TNFα) secretion, and a 67% reduction in interleukin-10 (IL-10) secretion when compared to LPS-stimulated control cells (P ≤ 0.01 for both). LPS did not stimulate TNFα or IL-10 secretion in adenovirus-mediated control or MyD88 knockdown cells. These data demonstrate that Raw 264.7 macrophages maintain responsiveness to inflammatory stimuli following CRISPR/Cas9-mediated reductions in MyD88, but not following adenovirus-mediated MyD88 knockdown.

Galactosemias are a family of autosomal recessive genetic disorders resulting from impaired enzymes of the Leloir pathway of galactose metabolism including galactokinase, galactose uridyltransferase, and UDP-galactose 4-epimerase that are critical for conversion of galactose into glucose-6-phosphate. To better understand pathophysiological mechanisms involved in galactosemia and develop novel therapies to address the unmet need in patients, it is important to develop reliable assays to measure the activity of the Leloir pathway enzymes. Here we describe in-depth methods for indirectly measuring galacose-1-phosphate uridyltransferase activity in cell culture and animal tissues.

The prevalence of kidney dysfunction continues to increase worldwide, driving the need to develop transplantable renal tissues. The kidney develops from four major renal progenitor populations: nephron epithelial, ureteric epithelial, interstitial and endothelial progenitors. Methods have been developed to generate kidney organoids but few or dispersed tubular clusters within the organoids hamper its use in regenerative applications. Here, we describe a detailed protocol of asynchronous mixing of kidney progenitors using organotypic culture conditions to generate kidney organoids tightly packed with tubular clusters and major renal structures including endothelial network and functional proximal tubules. This protocol provides guidance in the culture of human embryonic stem cells from a National Institute of Health-approved line and their directed differentiation into kidney organoids. Our 18-day protocol provides a rapid method to generate kidney organoids that facilitate the study of different nephrological events including in vitro tissue development, disease modeling and chemical screening. However, further studies are required to optimize the protocol to generate additional renal-specific cell types, interconnected nephron segments and physiologically functional renal tissues.

We describe a procedure of performing in silico (virtual) screening using a web-based service, the MTiOpenScreen, which is freely accessible to non-commercial users. We shall use the SARS-CoV-2 main protease as an example. Starting from a structure downloaded from the Protein Data Bank, we discuss how to prepare the coordinates file, taking into account the known biochemical background information of the target protein. The reader will find that this preparation step takes up most of the effort before the target is ready for screening. The steps for uploading the target structure and defining the search volume by critical residues, and the main parameters to use, are outlined. When this protocol is followed, the user will expect to obtain a ranked list of small approved drug compounds docked into the target structure. The results can be readily examined graphically on the web site or downloaded for studying in a local molecular graphics program such as PyMOL.

In vitro fermentation systems offer significant opportunity for deconvoluting complex metabolic dynamics within polymicrobial communities, particularly those associated with the human gut microbiome. In vitro gut models have broad experimental capacity allowing rapid evaluation of multiple parameters, generating knowledge to inform design of subsequent in vivo studies. Here, our method describes an in vitro fermentation test bed to provide a physiologically-relevant assessment of engineered probiotics circuit design functions. Typically, engineered probiotics are evaluated under pristine, mono- or co-culture conditions and transitioned directly into animal or human studies, commonly resulting in a loss of desired function when introduced to complex gut communities. Our method encompasses a systematic workflow entailing fermentation, molecular and functional characterization, and statistical analyses to validate an engineered probiotic's persistence, plasmid stability and reporter response. To demonstrate the workflow, simplified polymicrobial communities of human gut microbial commensals were utilized to investigate the probiotic Escherichia coli Nissle 1917 engineered to produce a fluorescent reporter protein. Commensals were assembled with increasing complexity to produce a mock community based on nutrient utilization. The method assesses engineered probiotic persistence in a competitive growth environment, reporter production and function, effect of engineering on organism growth and influence on commensal composition. The in vitro test bed represents a new element within the Design-Build-Test-Learn paradigm, providing physiologically-relevant feedback for circuit re-design and experimental validation for transition of engineered probiotics to higher fidelity animal or human studies.

Three-dimensional (3D) printing of biodegradable polymers has rapidly become a popular approach to create scaffolds for tissue engineering. This technique enables fabrication of complex architectures and layer-by-layer spatial control of multiple components with high resolution. The resulting scaffolds can also present distinct chemical groups or bioactive cues on the surface to guide cell behavior. However, surface functionalization often includes one or more post-fabrication processing steps, which typically produce biomaterials with homogeneously distributed chemistries that fail to mimic the biochemical organization found in native tissues. As an alternative, our laboratory developed a novel method that combines solvent-cast 3D printing with peptide-polymer conjugates to spatially present multiple biochemical cues in a single scaffold without requiring post-fabrication modification. Here, we describe a detailed, stepwise protocol to fabricate peptide-functionalized scaffolds and characterize their physical architecture and biochemical spatial organization. We used these 3D-printed scaffolds to direct human mesenchymal stem cell differentiation and osteochondral tissue formation by controlling the spatial presentation of cartilage-promoting and bone-promoting peptides. This protocol also describes how to seed scaffolds and evaluate matrix deposition driven by peptide organization.

Single-molecule studies involving DNA or RNA, require homogeneous preparations of nucleic acid substrates of exceptional quality. Over the past several years, a variety of methods have been published describing different purification methods but these are frustratingly inconsistent with variable yields even in the hands of experienced bench scientists. To address these issues, we present an optimized and straightforward, column-based approach that is reproducible and produces high yields of substrates or substrate components of exceptional quality. Central to the success of the method presented is the use of a non-porous anion exchange resin. In addition to the use of this resin, we encourage the optimization of each step in the construction of substrates. The fully optimized method produces high yields of a hairpin DNA substrate of exceptional quality. While this substrate is suitable for single-molecule, magnetic tweezer experiments, the described method is readily adaptable to the production of DNA substrates for the majority of single-molecule studies involving nucleic acids ranging in size from 70-15000 bp.

We describe a visual stimulus that can be used with both larval and adult zebrafish (Danio rerio). This protocol is a modification of a standard visual behavior analysis, the optomotor response (OMR). The OMR is often used to determine the spatial response or to detect directional visuomotor deficiencies. An OMR can be generated using a high contrast grated pattern, typically vertical bars. The spatial sensitivity is measured by detection and response to a change in grating bar width and is reported in cycles per degree (CPD). This test has been used extensively with zebrafish larvae and adults to identify visual- and/or motor-based mutations. Historically, when tested in adults, the grated pattern was presented from a vertical perspective, using a rotating cylinder around a holding tank, allowing the grating to be seen solely from the sides and front of the organism. In contrast, OMRs in zebrafish larvae are elicited using a stimulus projected below the fish. This difference in methodology means that two different experimental set-ups are required: one for adults and one for larvae. Our visual stimulus modifies the stimulation format so that a single OMR stimulus, suitable for use with both adults and larvae, is being presented underneath the fish. Analysis of visuomotor responses using this method does not require costly behavioral tracking software and, using a single behavioral paradigm, allows the observer to rapidly determine visual spatial response in both zebrafish larvae and adults.