Biochemistry

Progress in bioinformatics has facilitated the identification of a large number of putative glycosyltransferases (GTs) associated with many physiological processes. However, many of these GTs remain with unknown biochemical function due to numerous technical limitations. One of these limitations is the lack of innovative tools for large-scale screening of enzyme activity in vitro and testing protein–protein interactions (PPIs) between GT partners. Currently, testing the enzyme activity of a protein requires its production in a heterologous expression system and purification before enzyme assays, a process that is time-consuming and not amenable to high-throughput screening. To overcome this, we developed a platform called in vitro GT-array (i-GT-ray). In this platform, 96-well microplates are coated with plasmid DNA encoding for tagged GTs and a capture antibody. Tagged GTs are produced from plasmid DNA via a cell-free in vitro transcription/translation (IVTT) system and captured through the anti-tag capture antibody directly on microplates. After washing to remove IVTT components, the captured enzymes can be considered purified, and their activity can be tested directly on microplates. The whole process can be performed in less than two days, compared to several weeks for currently available screening methods. The i-GT-ray platform has also been adapted to investigate PPIs between GTs. Here, we provide a practical user guide for the preparation of GT-arrays coated with plasmid DNA and a capture antibody that can be used for monitoring enzyme activity and PPIs of GTs in a high-throughput manner.

Accurate quantification of von Willebrand factor ristocetin cofactor activity (VWF:RCo) is critical for the diagnosis and classification of von Willebrand disease, the most common hereditary and acquired bleeding disorder in humans. Moreover, it is important to accurately assess the function of von Willebrand factor (VWF) concentrates within the pharmaceutical industry to provide consistent and high-quality biopharmaceuticals. Although the performance of VWF:RCo assay has been improved by using coagulation analyzers, which are specialized devices for blood and blood plasma samples, scientists still report a high degree of intra- and inter-assay variation in clinical laboratories. Moreover, high, manual sample dilutions are required for VWF:RCo determination of VWF concentrates within the pharmaceutical industry, which are a major source for assay imprecision. For the first time, we present a precise and accurate method to determine VWF:RCo, where all critical pipetting and mixing steps are automated. A pre-dilution setup was established on CyBio FeliX (Analytik-Jena) liquid handling system, and an adapted VWF:RCo method on BCS-XP analyzer (Siemens) is used. The automated pre-dilution method was executed on three different, most frequently used coagulation analyzers and compared to manual pre-dilutions performed by an experienced operator. Comparative sample testing revealed a similar assay precision (coefficient of variation = 5.9% automated, 3.1% manual pre-dilution) and no significant differences between the automated approach and manual dilutions of an expert in this method. While no outliers were generated with the automated procedure, the manual pre-dilution resulted in an error rate of 8.3%. Overall, this operator-independent protocol enables standardization and offers an efficient way of fully automating VWF activity assays, while maintaining the precision and accuracy of an expert analyst.

Generating protein conjugates using the bioorthogonal ligation between tetrazines and trans-cyclooctene groups avoids the need to manipulate cysteine amino acids; this ligation is rapid, site-specific, and stoichiometric and allows for labeling of proteins in complex biological environments. Here, we provide a protocol for the expression of conjugation-ready proteins at high yields in Escherichia coli with greater than 95% encoding and labeling fidelity. This protocol focuses on installing the Tet2 tetrazine amino acid using an optimized genetic code expansion (GCE) machinery system, Tet2 pAJE-E7, to direct Tet2 encoding at TAG stop codons in BL21 E. coli strains, enabling reproducible expression of Tet2-proteins that quantitatively react with trans-cyclooctene (TCO) groups within 5 min at room temperature and physiological pH. The use of the BL21 derivative B95(DE3) minimizes premature truncation byproducts caused by incomplete suppression of TAG stop codons, which makes it possible to use more diverse protein construct designs. Here, using a superfolder green fluorescent protein construct as an example protein, we describe in detail a four-day process for encoding Tet2 with yields of ~200 mg per liter of culture. Additionally, a simple and fast diagnostic gel electrophoretic mobility shift assay is described to confirm Tet2-Et encoding and reactivity. Finally, strategies are discussed to adapt the protocol to alternative proteins of interest and optimize expression yields and reactivity for that protein.

Bottom-up proteomics utilizes sample preparation techniques to enzymatically digest proteins, thereby generating identifiable and quantifiable peptides. Proteomics integrates with other omics methodologies, such as genomics and transcriptomics, to elucidate biomarkers associated with diseases and responses to drug or biologics treatment. The methodologies employed for preparing proteomic samples for mass spectrometry analysis exhibit variability across several factors, including the composition of lysis buffer detergents, homogenization techniques, protein extraction and precipitation methodologies, alkylation strategies, and the selection of digestion enzymes. The general workflow for bottom-up proteomics consists of sample preparation, mass spectrometric data acquisition (LC-MS/MS analysis), and subsequent downstream data analysis including protein quantification and differential expression analysis. Sample preparation poses a persistent challenge due to issues such as low reproducibility and inherent procedure complexities. Herein, we have developed a validated chloroform/methanol sample preparation protocol to obtain reproducible peptide mixtures from both rodent tissue and human cell line samples for bottom-up proteomics analysis. The protocol we established may facilitate the standardization of bottom-up proteomics workflows, thereby enhancing the acquisition of reliable biologically and/or clinically relevant proteomic data.

Microscale thermophoresis (MST) is a technique used to measure the strength of molecular interactions. MST is a thermophoretic-based technique that monitors the change in fluorescence associated with the movement of fluorescent-labeled molecules in response to a temperature gradient triggered by an IR LASER. MST has advantages over other approaches for examining molecular interactions, such as isothermal titration calorimetry, nuclear magnetic resonance, biolayer interferometry, and surface plasmon resonance, requiring a small sample size that does not need to be immobilized and a high-sensitivity fluorescence detection. In addition, since the approach involves the loading of samples into capillaries that can be easily sealed, it can be adapted to analyze oxygen-sensitive samples. In this Bio-protocol, we describe the troubleshooting and optimization we have done to enable the use of MST to examine protein–protein interactions, protein–ligand interactions, and protein–nanocrystal interactions. The salient elements in the developed procedures include 1) loading and sealing capabilities in an anaerobic chamber for analysis using a NanoTemper MST located on the benchtop in air, 2) identification of the optimal reducing agents compatible with data acquisition with effective protection against trace oxygen, and 3) the optimization of data acquisition and analysis procedures. The procedures lay the groundwork to define the determinants of molecular interactions in these technically demanding systems.

Efficient and nontoxic delivery of foreign cargo into cells is a critical step in many biological studies and cell engineering workflows with applications in areas such as biomanufacturing and cell-based therapeutics. However, effective molecular delivery into cells involves optimizing several experimental parameters. In the case of electroporation-based intracellular delivery, there is a need to optimize parameters like pulse voltage, duration, buffer type, and cargo concentration for each unique application. Here, we present the protocol for fabricating and utilizing a high-throughput multi-well localized electroporation device (LEPD) assisted by deep learning–based image analysis to enable rapid optimization of experimental parameters for efficient and nontoxic molecular delivery into cells. The LEPD and the optimization workflow presented herein are relevant to both adherent and suspended cell types and different molecular cargo (DNA, RNA, and proteins). The workflow enables multiplexed combinatorial experiments and can be adapted to cell engineering applications requiring in vitro delivery.

Peripheral membrane proteins (PMPs) are a subgroup of membrane-associated proteins that are water-soluble and bind to membranes, often reversibly, to perform their function. These proteins have been extensively studied in the aqueous state, but there is often a lack of high-resolution structural and functional studies of these proteins in the membrane-bound state. Currently, nuclear magnetic resonance (NMR) is among the best-equipped methods to study these relatively small proteins and domains, but current models have some disadvantages that prevent a full understanding of PMP interactions with membranes and lipids. Micelles, bicelles, and nanodiscs are all available for NMR observation but are based on synthetic lipids that may destabilize proteins or are too large to accommodate straightforward structural analysis. This protocol introduces a method for forming reverse micelles using lipids from natural sources, here called native reverse micelles. This technique allows the PMPs to embed within a shell of naturally derived lipids surrounding a small water core solubilized in an alkane solvent. PMP embedment in the lipid shell mimics binding to a cellular membrane. Here, naturally derived lipids from soy, bovine heart, and porcine brain are used in conjunction with n-dodecylphosphocholine (DPC) to encapsulate a PMP from either concentrated or dried protein, resulting in reverse micelles that may be confirmed via dynamic light scattering and NMR. This protocol allows for high-quality NMR data of PMPs interacting with membrane lipids within a biologically accurate environment.

Microglia, the brain's primary resident immune cell, exists in various phenotypic states depending on intrinsic and extrinsic signaling. Distinguishing between these phenotypes can offer valuable biological insights into neurodevelopmental and neurodegenerative processes. Recent advances in single-cell transcriptomic profiling have allowed for increased granularity and better separation of distinct microglial states. While techniques such as immunofluorescence and single-cell RNA sequencing (scRNA-seq) are available to differentiate microglial phenotypes and functions, these methods present notable limitations, including challenging quantification methods, high cost, and advanced analytical techniques. This protocol addresses these limitations by presenting an optimized cell preparation procedure that prevents ex vivo activation and a flow cytometry panel to distinguish four distinct microglial states from murine brain tissue. Following cell preparation, fluorescent antibodies were applied to label 1) homeostatic, 2) disease-associated (DAM), 3) interferon response (IRM), and 4) lipid-droplet accumulating (LDAM) microglia, based on gene markers identified in previous scRNA-seq studies. Stained cells were analyzed by flow cytometry to assess phenotypic distribution as a function of age and sex. A key advantage of this procedure is its adaptability, allowing the panel provided to be enhanced using additional markers with an appropriate cell analyzer (i.e., Cytek Aurora 5 laser spectral flow cytometer) and interrogating different brain regions or disease models. Additionally, this protocol does not require microglial cell sorting, resulting in a relatively quick and straightforward experiment. Ultimately, this protocol can compare the distribution of microglial phenotypic states between various experimental groups, such as disease state or age, with a lower cost and higher throughput than scRNA-seq.

The Auxin-inducible degron (AID) system is a genetic tool that induces rapid target protein depletion in an auxin-dependent manner. Recently, two advanced AID systems—the super-sensitive AID and AID 2—were developed using an improved pair of synthetic auxins and mutated TIR1 proteins. In these AID systems, a nanomolar concentration of synthetic auxins is sufficient as a degradation inducer for target proteins. However, despite these advancements, AID systems still require the fusion of an AID tag to the target protein for degradation, potentially affecting its function and stability. To address this limitation, we developed an affinity linker–based super-sensitive AID (AlissAID) system using a single peptide antibody known as a nanobody. In this system, the degradation of GFP- or mCherry-tagged target proteins is induced in a synthetic auxin (5-Ad-IAA)–dependent manner. Here, we introduce a simple method for generating AlissAID strains targeting GFP or mCherry fusion proteins in budding yeasts.

Agrobacterium-mediated transient gene expression in Nicotiana benthamiana is widely used to study gene function in plants. One dramatic phenotype that is frequently screened for is cell death. Here, we present a simplified protocol for Agrobacterium-mediated transient gene expression by infiltration. Compared with current methods, the novel protocol can be done without a centrifuge or spectrometer, thereby suitable for K-12 outreach programs as well as rapidly identifying genes that induce cell death.