Biochemistry

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.

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).

Palmitoylation is a unique and reversible posttranslational lipid modification (PTM) that plays a critical role in many cellular events, including protein stability, activity, membrane association, and protein–protein interactions. The dynamic nature of palmitoylation dictates the efficient sorting of various retinal proteins to specific subcellular compartments. However, the underlying mechanism through which palmitoylation supports efficient protein trafficking in the retina remains unclear. Recent studies show that palmitoylation can also function as a signaling PTM, underlying epigenetic regulation and homeostasis in the retina. Efficient isolation of retinal palmitoyl proteome will pave the way to a better understanding of the role(s) for palmitoylation in visual function. The standard methods for detecting palmitoylated proteins employ ³H- or ¹⁴C-radiolabeled palmitic acid and have many limitations, including poor sensitivity. Relatively recent studies use thiopropyl Sepharose 6B resin, which offers efficient detection of palmitoylated proteome but is now discontinued from the market. Here, we describe a modified acyl resin–assisted capture (Acyl-RAC) method using agarose S3 high-capacity resin to purify palmitoylated proteins from the retina and other tissues, which is greatly compatible with downstream processing by LC-MS/MS. Unlike other palmitoylation assays, the present protocol is easy to perform and cost-effective.

Graphical overview

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.