Histone acetyltransferases (HATs, also known as lysine acetyltransferases, KATs) catalyze acetylation of their cognate protein substrates using acetyl‐CoA (Ac‐CoA) as a cofactor and are involved in various physiological and pathological processes. Advances in mass spectrometry‐based proteomics have allowed the discovery of thousands of acetylated proteins and the specific acetylated lysine sites. However, due to the rapid dynamics and functional redundancy of HAT activities, and the limitation of using antibodies to capture acetylated lysines, it is challenging to systematically and precisely define both the substrates and sites directly acetylated by a given HAT. Here, we describe a chemoproteomic approach to identify and profile protein substrates of individual HAT enzymes on the proteomic scale. The approach involves protein engineering to enlarge the Ac‐CoA binding pocket of the HAT of interest, such that a mutant form is generated that can use functionalized acyl‐CoAs as a cofactor surrogate to bioorthogonally label its protein substrates. The acylated protein substrates can then be chemoselectively conjugated either with a fluorescent probe (for imaging detection) or with a biotin handle (for streptavidin pulldown and chemoproteomic identification). This modular chemical biology approach has been successfully implemented to identify protein substrates of p300, GCN5, and HAT1, and it is expected that this method can be applied to profile and identify the sub‐acetylomes of many other HAT enzymes. © 2022 Wiley Periodicals LLC.
Protein S‐acylation, predominately in the form of palmitoylation, is a reversible lipid post‐translational modification on cysteines that plays important roles in protein localization, trafficking, activity, and complex assembly. The functions and regulatory mechanisms of S‐acylation have been extensively studied in mammals owing to remarkable development of high‐resolution proteomics and the discovery of the S‐acylation‐related enzymes. However, the advancement of S‐acylation studies in plants lags behind that in mammals, mainly due to the lack of knowledge about proteins responsible for this process, such as protein acyltransferases and their substrates. In this article, a set of systematic protocols to study global S‐acylation in
- NSF-PAR ID:
- 10238887
- Publisher / Repository:
- Wiley Blackwell (John Wiley & Sons)
- Date Published:
- Journal Name:
- Current Protocols in Plant Biology
- Volume:
- 5
- Issue:
- 4
- ISSN:
- 2379-8068
- Format(s):
- Medium: X
- Sponsoring Org:
- National Science Foundation
More Like this
-
Abstract Basic Protocol 1 : Labeling HAT protein substrates with azide/alkyne‐biotinAlternate Protocol : Labeling protein substrates of HATs with azide/alkyne‐TAMRA for in‐gel visualizationSupport Protocol 1 : Expression and purification of HAT mutantsSupport Protocol 2 : Synthesis of Ac‐CoA surrogatesBasic Protocol 2 : Streptavidin enrichment of biotinylated HAT substratesBasic Protocol 3 : Chemoproteomic identification of HAT substratesBasic Protocol 4 : Validation of specific HAT substrates with western blotting -
Abstract Histone post‐translational modifications (PTMs) play important roles in many biological processes, including gene regulation and chromatin dynamics, and are thus of high interest across many fields of biological research. Chromatin immunoprecipitation coupled with sequencing (ChIP‐seq) is a powerful tool to profile histone PTMs
in vivo . This method, however, is largely dependent on the specificity and availability of suitable commercial antibodies. While mass spectrometry (MS)–based proteomic approaches to quantitatively measure histone PTMs have been developed in mammals and several other model organisms, such methods are currently not readily available in plants. One major challenge for the implementation of such methods in plants has been the difficulty in isolating sufficient amounts of pure, high‐quality histones, a step rendered difficult by the presence of the cell wall. Here, we developed a high‐yielding histone extraction and purification method optimized forArabidopsis thaliana that can be used to obtain high‐quality histones for MS. In contrast to other methods used in plants, this approach is relatively simple, and does not require membranes or additional specialized steps, such as gel excision or chromatography, to extract highly purified histones. We also describe methods for producing MS‐ready histone peptides through chemical labeling and digestion. Finally, we describe an optimized method to quantify and analyze the resulting histone PTM data using a modified version of EpiProfile 2.0 for Arabidopsis. In all, the workflow described here can be used to measure changes to histone PTMs resulting from various treatments, stresses, and time courses, as well as in different mutant lines. © 2022 Wiley Periodicals LLC.Basic Protocol 1 : Nuclear isolation and histone acid extractionBasic Protocol 2 : Peptide labeling, digestion, and desaltingBasic Protocol 3 : Histone HPLC‐MS/MS and data analysis -
Abstract Extracellular vesicles (EVs) in plants have emerged as key players in cell‐to‐cell communication and cross‐kingdom RNAi between plants and pathogens by facilitating the exchange of RNA, proteins, and other molecules. In addition to their role in intercellular communication, plant EVs also show promise as potential therapeutics and indicators of plant health. However, plant EVs exhibit significant heterogeneity in their protein markers, size, and biogenesis pathways, strongly influencing their composition and functionality. While mammalian EVs can be generally classified as exosomes that are derived from multivesicular bodies (MVBs), microvesicles that are shed from the plasma membrane, or as apoptotic bodies that originate from cells undergoing apoptosis, plant EVs remain poorly studied in comparison. At least three subclasses of EVs have been identified in
Arabidopsis leaves to date, including Tetraspanin‐positive exosomes derived from MVBs, Penetration 1 (PEN1)‐positive EVs, and EVs derived from exocyst‐positive organelles (EXPO). Differences in the plant starting material and isolation techniques have resulted in different purities, quality, and compositions of the resulting EVs, complicating efforts to better understand the role of these EVs in plants. We performed a comparative analysis on commonly used plant EV isolation methods and have identified an effective protocol for extracting clean apoplastic washing fluid (AWF) and isolating high‐quality intact and pure EVs ofArabidopsis thaliana . These EVs can then be used for various applications or studied to assess their cargos and functionality in plants. Furthermore, this process can be easily adapted to other plant species of interest. © 2022 Wiley Periodicals LLC.Basic Protocol 1 : Isolation of EVs from the apoplastic fluid ofArabidopsis thaliana Basic Protocol 2 : Density gradient fractionation of EVsBasic Protocol 3 : Immuno‐isolation of EVs usingArabidopsis tetraspanin 8 (TET8) antibody -
Abstract Numerous methods have been developed in model systems to deplete or inactivate proteins to elucidate their functional roles. In
Caenorhabditis elegans , a common method for protein depletion is RNA interference (RNAi), in which mRNA is targeted for degradation.C. elegans is also a powerful genetic organism, amenable to large‐scale genetic screens and CRISPR‐mediated genome editing. However, these approaches largely lead to constitutive inhibition, which can make it difficult to study proteins essential for development or to dissect dynamic cellular processes. Thus, there have been recent efforts to develop methods to rapidly inactivate or deplete proteins to overcome these barriers. One such method that is proving to be exceptionally powerful is auxin‐inducible degradation. In order to apply this approach inC. elegans , a 44–amino acid degron tag is added to the protein of interest, and theArabidopsis ubiquitin ligase TIR1 is expressed in target tissues. When the plant hormone auxin is added, it mediates an interaction between TIR1 and the degron‐tagged protein of interest, which triggers ubiquitination of the protein and its rapid degradation via the proteasome. Here, we have outlined multiple methods for inducing auxin‐mediated depletion of target proteins inC. elegans , highlighting the versatility and power of this method. © 2021 Wiley Periodicals LLC.This article was corrected on 19 July 2022. See the end of the full text for details.
Basic Protocol 1 : Long‐term auxin‐mediated depletion on platesSupport Protocol : Preparation of NGM and NGM‐auxin platesBasic Protocol 2 : Rapid auxin‐mediated depletion via soakingBasic Protocol 3 : Acute auxin‐mediated depletion in isolated embryosBasic Protocol 4 : Assessing auxin‐mediated depletion -
Abstract Class II major histocompatibility complex peptide (MHC‐IIp) multimers are precisely engineered reagents used to detect T cells specific for antigens from pathogens, tumors, and self‐proteins. While the related Class I MHC/peptide (MHC‐Ip) multimers are usually produced from subunits expressed in
E. coli , most Class II MHC alleles cannot be produced in bacteria, and this has contributed to the perception that MHC‐IIp reagents are harder to produce. Herein, we present a robust constitutive expression system for soluble biotinylated MHC‐IIp proteins that uses stable lentiviral vector−transduced derivatives of HEK‐293T cells. The expression design includes allele‐specific peptide ligands tethered to the amino‐terminus of the MHC‐II β chain via a protease‐cleavable linker. Following cleavage of the linker, HLA‐DM is used to catalyze efficient peptide exchange, enabling high‐throughput production of many distinct MHC‐IIp complexes from a single production cell line. Peptide exchange is monitored using either of two label‐free methods, native isoelectric focusing gel electrophoresis or matrix‐assisted laser desorption/ionization time‐of‐flight (MALDI‐TOF) mass spectrometry of eluted peptides. Together, these methods produce MHC‐IIp complexes that are highly homogeneous and that form the basis for excellent MHC‐IIp multimer reagents. © 2021 Wiley Periodicals LLC.This article was corrected on 19 July 2022. See the end of the full text for details.
Basic Protocol 1 : Lentivirus production and expression line creationSupport Protocol 1 : Six‐well assay for estimation of production cell line yieldSupport Protocol 2 : Universal ELISA for quantifying proteins with fused leucine zippers and His‐tagsBasic Protocol 2 : Cultures for production of Class II MHC proteinsBasic Protocol 3 : Purification of Class II MHC proteins by anti‐leucine zipper affinity chromatographyAlternate Protocol 1 : IMAC purification of His‐tagged Class II MHCSupport Protocol 3 : Protein concentration measurements and adjustmentsSupport Protocol 4 : Polishing purification by anion‐exchange chromatographySupport Protocol 5 : Estimating biotinylation percentage by streptavidin precipitationBasic Protocol 4 : Peptide exchangeBasic Protocol 5 : Analysis of peptide exchange by matrix‐assisted laser desorption/ionization (MALDI) mass spectrometryAlternate Protocol 2 : Native isoelectric focusing to validate MHC‐II peptide loadingBasic Protocol 6 : MultimerizationBasic Protocol 7 : Staining cells with Class II MHC tetramers