生物化学

Protein O-GlcNAcylation is a prevalent and dynamic post-translational modification that targets a multitude of nuclear and cytoplasmic proteins. Through the modification of diverse substrates, O-GlcNAcylation plays a pivotal role in essential cellular processes, including transcription, translation, and protein homeostasis. Dysregulation of O-GlcNAc homeostasis has been implicated in a variety of diseases, including cardiovascular diseases, cancer, and neurodegenerative diseases. Studying O-GlcNAcylated proteins in different tissues is crucial to understanding the pathogenesis of these diseases. However, identifying phenotype-relevant candidate substrates in a tissue-specific manner remains unfeasible. We developed a novel tool for the analysis of O-GlcNAcylated proteins, combining a catalytically inactive CpOGA mutant CpOGA^CD and TurboID proximity labeling technology. This tool converts O-GlcNAc modifications into biotin labeling, enabling the enrichment and mass spectrometry (MS) identification of O-GlcNAcylated proteins in specific tissues. Meanwhile, TurboID-CpOGA^DM, which carries two point mutations that inactivate both its catalytic and binding activities toward O-GlcNAc modification, was used as a control to differentiate O-GlcNAc-independent protein–protein interactions. We have successfully used TurboID-CpOGA^CD/DM (TurboID-CpOGA^M) to enrich O-GlcNAc proteins in Drosophila combining the UAS/Gal4 system. Our protocol provides a comprehensive workflow for tissue-specific enrichment of candidate O-GlcNAcylated substrates and offers a valuable tool for dissecting tissue-specific O-GlcNAcylation functions in Drosophila.

Bioorthogonal chemical reporters are non-native chemical handles introduced into biomolecules of living systems, typically through metabolic or protein engineering. These functionalities can undergo bioorthogonal reactions, such as copper-catalyzed alkyne-azide cycloaddition (CuAAC), with small-molecule probes, enabling the tagging and visualization of biomolecules. This approach has greatly enhanced our understanding of cellular dynamics, enzyme targeting, and protein post-translational modifications. Herein, we report a protocol for preparing protein lysates for click reaction and in-gel fluorescence analysis, exemplified using alk-16, a terminal alkyne-functionalized stearic acid analog, to investigate proteins with fatty acylation. This protocol provides methods for the fluorescent visualization of chemical reporter–labeled proteomes or individual proteins of interest (POIs).

This protocol describes the recombinant expression of proteins in E. coli containing phosphoserine (pSer) installed at positions guided by TAG codons. The E. coli strains that can be used here are engineered with a ∆serB genomic knockout to produce pSer internally at high levels, so no exogenously added pSer is required, and the addition of pSer to the media will not affect expression yields. For “truncation-free” expression and improved yields with high flexibility of construct design, it is preferred to use the Release Factor-1 (RF1) deficient strain B95(DE3) ∆A ∆fabR ∆serB, though use of the standard RF1-containing BL21(DE3) ∆serB is also described. Both of these strains are serine auxotrophs and will not grow in standard minimal media. This protocol uses rich auto-induction media for streamlined and maximal production of homogeneously modified protein, yielding ~100–200 mg of single pSer-containing sfGFP per liter of culture. Using this genetic code expansion (GCE) approach, in which pSer is installed into proteins during translation, allows researchers to produce milligram quantities of specific phospho-proteins without requiring kinases, which can be purified for downstream in vitro studies relating to phosphorylation-dependent signaling systems, protein regulation by phosphorylation, and protein–protein interactions.

Graphical abstract:

Cortactin is an actin-binding protein that regulates processes like cell migration, endocytosis, and tumor cell metastasis. Although cortactin is associated with actin-cytoskeletal dynamics in non-neuronal cells and cell-free systems, the exact mechanisms underlying its fundamental roles in neuronal growth cones are not fully explored. Recent reports show that Aplysia Src2 tyrosine kinase induces phosphorylation of cortactin as a mechanism to control lamellipodia protrusion and filopodia formation in cultured Aplysia bag cell neurons (He et al., 2015; Ren et al., 2019). In order to provide in vitro evidence for Src2-mediated phosphorylation of cortactin, we developed a robust and cost-effective method for the efficient expression and purification of Aplysia cortactin and Src2 kinase that can be used for biochemical studies including phosphorylation assays. By co-purifying cortactin and Src kinase with a phosphatase (YopH) from Yersinia enterocolitica, we eliminated the problem of non-specific phosphorylation of induced proteins by bacterial kinases and also reduced costs by bypassing the need for commercial enzymatic treatments. This protocol is reproducible and can be modified to produce homogenous non-phosphorylated proteins during recombinant protein expression in Escherichia coli.

Mitochondria are essential organelles containing approximately 1,500 proteins. Only approximately 1% of these proteins are synthesized inside mitochondria, whereas the remaining 99% are synthesized as precursors on cytosolic ribosomes and imported into the organelle. Various tools and techniques to analyze the import process have been developed. Among them, in vitro reconstituted import systems are of importance to study these processes in detail. These experiments monitor the import reaction of mitochondrial precursors that were previously radiolabeled in a cell-free environment. However, the methods described have been mostly performed in mitochondria isolated from S. cerevisiae. Here, we describe the adaptation of this powerful assay to import proteins into crude mitochondria isolated from human tissue culture cells.

Graphic abstract:

Overview of the assay to monitor protein import into mitochondria isolated from human cells

Hydrogen sulfide (H₂S) is emerging as an important modulator in bacterial cytoprotection against the host immune response in infected animals, which may well be attributed to downstream highly oxidized sulfur species, termed reactive sulfur species (RSS), derived from H₂S. One mechanism by which H₂S/RSS may signal in the cell is through proteome S-sulfuration (persulfidation), which is the conversion of protein thiols (-SH) to protein persulfides (-SSH). While several analytical methods have been developed to profile sites of protein persulfidation, few have been applied to bacterial cells. The analytical workflow presented here was recently utilized to profile proteome persulfidation in the major human pathogen Acinetobacter baumannii treated with an exogenous sulfide source, Na₂S. The data obtained using this protocol allow quantitation of the change in persulfidation status of each cysteine in the proteome normalized to the change in protein abundance, thus identifying sites of persulfidation that may constitute regulatory modifications. These can be validated using follow-up biochemical studies.

Palmitoylation refers to the modification of the cysteine thiols in proteins by fatty acids, most commonly palmitic acid, through ‘thioester bond’ formation. In vivo, palmitoylation of proteins is catalyzed by palmitoyl acyltransferases (PATs or DHHC-PATs). Palmitoylation has recently emerged as a crucial post-translational modification in malarial parasites. The expression and activity of palmitoyl transferases vary across different developmental stages of the malarial parasite’s life cycle. The abundance of palmitoylated proteins at a given stage is a measure of overall PAT activity. The PAT activity can also change in response to external signals or inhibitors. Here, we describe a protocol to ‘image’ palmitoyl-transferase activity during the asexual stages using Click Chemistry and fluorescence microscopy. This method is based on metabolic labeling of a clickable analog of palmitic acid by parasitic cells, followed by CuAAC (Copper-catalyzed Alkyne-Azide Cycloaddition reaction) Click Chemistry to render palmitoylated proteins fluorescent. Fluorescence allows the quantitation of intracellular palmitoylation in parasite cells across various development stages. Using this method, we observed that intracellular palmitoylation increases as the parasite transitions from ring to schizont stages and appears to be most abundant during the schizont stages in Plasmodium falciparum.

Glutamylation is a posttranslational modification where the amino group of a free glutamate amino acid is conjugated to the carboxyl group of a glutamate side chain within a target protein. SidJ is a Legionella kinase-like protein that has recently been identified to perform protein polyglutamylation of the Legionella SdeA Phosphoribosyl-Ubiquitin (PR-Ub) ligase to inhibit SdeA’s activity. The attachment of multiple glutamate amino acids to the catalytic glutamate residue of SdeA by SidJ inhibits SdeA’s modification of ubiquitin (Ub) and ligation activity. In this protocol, we will discuss a SidJ non-radioactive, in vitro glutamylation assay using its substrate SdeA. This will also include a second reaction to assay the inhibition of SdeA by using both modification of free Ub and ligation of ADP-ribosylated Ubiquitin (ADPR-Ub) to SdeA’s substrate Rab33b. Prior to the identification and publication of SdeA’s activity, no SdeA inhibition assays existed. Our group and others have demonstrated various methods to display inhibition of SdeA’s activity. The alternatives include measurement of ADP-ribosylation of Ub using radioactive NAD, NAD hydrolysis, and Western blot analysis of HA-Ub ligation by SdeA. This protocol will describe the inhibition of both ubiquitin modification and the PR-Ub ligation by SdeA using inexpensive standard gels and Coomassie staining.

The Legionella effector protein SidJ has recently been identified to perform polyglutamylation on another Legionella effector, SdeA, ablating SdeA’s activity. SidJ is a kinase-like protein that requires the small eukaryotic protein calmodulin to perform glutamylation. Glutamylation is a relatively uncommon type of post-translational modification, where the amino group of a free glutamate amino acid is covalently linked to the γ-carboxyl group of a glutamate sidechain in a substrate protein. This protocol describes the SidJ glutamylation reaction using radioactive [U-¹⁴C] glutamate and its substrate SdeA, the separation of proteins by gel electrophoresis, preparation of gels for radioactive exposure, and relative quantification of glutamylation activity. This procedure is useful for the identification of substrates for glutamylation, characterization of substrate and glutamylase activities due to mutations, and identification of proteins with glutamylation activity. Some studies have assayed glutamylation with the use of [³H] glutamate (Regnard et al., 1998) and the use of the GT335 antibody (Wolff et al., 1992). However, the use of [U-¹⁴C] glutamate requires a shorter radioactive exposure time with no dependence on antibody specificity.

Chromatin modifications, like histone post translational modifications (PTMs), are critical for tuning gene expression and many other aspects of cell phenotype. Liquid chromatography coupled to mass spectrometry (LC-MS) has become the most suitable method to analyze histones and histone PTMs in a large-scale manner. Selected histone PTMs have known functions, and their aberrant regulation is linked to a wide variety of diseases, including cancer. However, histone analysis is scarcely used in diagnostics, partially due to the limited throughput and not ideal reproducibility of LC-MS based analysis. We describe a workflow that allows for high-throughput sample preparation is less than a day using 96-well plates. Following preparation, samples are sprayed into MS without LC, using an automated direct injection (DI-MS) method. Each analysis provides accurate quantification for 29 peptide sequences with 45 PTMs (methylations, acetylations and phosphorylations) for a total of 151 histone marks plus 16 unmodified histone peptides for relative quantification of histone variants. This workflow allows for < 1 min MS runs and higher reproducibility and robustness due to the absence of carryover or LC-based batch effects. Finally, we describe an engineered peptide sequence used to accurately monitor the efficiency of sample preparation, which can be detected during the DI-MS run.