Biochemistry

Small ubiquitin-related modifiers (SUMOs) are covalently conjugated onto the proteome and serve as signaling molecules in many aspects of eukaryotic cell biology, from S. cerevisiae and C. elegans to H. sapiens. The conjugatable SUMO variants, SUMO1 and the almost identical SUMO2 and SUMO3 (designated SUMO2/3), are processed by an E1(SAE1:SAE2)-E2(UBC9)-E3 enzyme cascade to produce SUMO-modified proteins. The prerogative of the SUMO biology field is to identify and study the specific proteins undergoing SUMOylation, which grants us insights into the biological pathway of interest. This protocol was developed using the human osteosarcoma cell line U2OS to enable the investigation of SUMO conjugates in mitosis, the cell division phase of the cell cycle. We enrich the cell population for mitotic cells, which are isolated and subjected to stringent lysis conditions involving a high concentration of SDS and DTT in RIPA buffer, to promote complete protein denaturation. The lysates in high SDS RIPA buffer are diluted to reduce the overall SDS concentration and undergo conventional immunoprecipitation using SUMO1- or SUMO2/3-specific antibodies bound to protein A/G agarose beads. The samples are then compatible with downstream readouts such as western blots and mass spectrometry. This protocol detects endogenous SUMOylated proteins and avoids exogenous SUMO overexpression, which can alter SUMO conjugate formation. Furthermore, this denaturing protocol ensures only SUMOylated proteins are immunoprecipitated, and not their interactors.

N-glycosylation is a ubiquitous post-translational modification (PTM) that regulates protein folding, stability, and biological function. Accurate identification and validation of N-glycosylation are therefore critical for understanding how glycosylation modulates protein activity. Here, we present a robust workflow for analyzing protein N-glycosylation in both animal and plant systems using peptide-N⁴-(N-acetyl-β-glucosaminyl) asparagine-amidase A and F (PNGase A and PNGase F). After enzymatic cleavage of the asparagine-linked N-glycans, samples are analyzed by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) and western blotting (WB) to detect shifts in apparent molecular weight (MW) indicative of deglycosylation. Key steps include denaturing the protein to expose glycosylation sites, optimizing buffer conditions for PNGase F and A treatment, and comparing glycosylated vs. deglycosylated forms by electrophoretic mobility. A troubleshooting guide addresses common challenges, including incomplete deglycosylation and low transfer efficiency during WB, offering practical solutions to ensure reliable results. This protocol provides researchers with a standardized, cost-effective framework for investigating protein N-glycosylation in diverse systems, from cell lysates to purified proteins, in both animal and plant models.

OtUBD is a high-affinity ubiquitin-binding domain (UBD) derived from a large protein produced by the microorganism Orientia tsutsugamushi. The following protocol describes a step-by-step process for the enrichment of ubiquitinated proteins from baker's yeast and mammalian cell lysates using OtUBD. The OtUBD affinity resin can strongly enrich both mono- and poly-ubiquitinated proteins from crude lysates. The protocol further describes the use of different buffer formulations to specifically enrich for proteins covalently modified by ubiquitin with or without proteins that associate with them. Combining different OtUBD-mediated enrichment protocols with liquid chromatography–tandem mass spectrometry (LC–MS/MS) helps distinguish the pool of covalently ubiquitinated proteins (the ubiquitinome) from ubiquitin- or ubiquitinated protein-interacting proteins (the ubiquitin interactome). The OtUBD tool described in the protocol has been used successfully with downstream applications such as immunoblotting and differential proteomics. It provides researchers with a versatile and economical tool for the study of ubiquitin biology.

The core planar cell polarity (PCP) protein Vang/Vangl, including Vangl1 and Vangl2 in vertebrates, is indispensable during development. Our previous studies showed that the activity of Vangl is tightly controlled by two important posttranslational modifications, ubiquitination and phosphorylation. Vangl is ubiquitinated through an endoplasmic reticulum-associated degradation (ERAD) pathway and is phosphorylated by casein kinase 1 (CK1) in response to Wnt. Here, we present step-by-step procedures to analyze Vangl ubiquitination and phosphorylation, including cell culture, transfection, sample preparation, and signal detection, as well as the use of newly available phospho-specific antibodies to detect Wnt-induced Vangl2 phosphorylation. The protocol described here can be applicable to the analysis of posttranslational modifications of other membrane proteins.

Understanding the folding pathway of any protein is of utmost importance for deciphering the folding problems under adverse conditions. We can obtain important information about the folding pathway by monitoring the folding of any protein from its unfolded state. It is usually very difficult to monitor the folding process in real time as the process is generally very fast, and we need a suitable read out. In this protocol, we have solved this issue by using a protein that is non-fluorescent in its unfolded state but fluoresces in its native state after folding. The kinetics of refolding can be monitored by following the increase in fluorescence in real time. Previously, this was generally achieved by either monitoring a protein’s enzymatic activity or measuring the tryptophan fluorescence, where the signal output depends on well-described enzymatic activity or the frequency of tryptophan residues present in the proteins, respectively. Here, we describe a simple and real-time assay to monitor the refolding of sGFP, a recently described slow-folding mutant of yeGFP (yeast enhanced GFP). We unfold this protein using chemical denaturant and refold in a suitable buffer, monitoring the increase in fluorescence over time. GFP is fluorescent only when correctly folded; thus, using this technique, we can measure the true rate of protein refolding by following the increase in fluorescence over time. Therefore, sGFP can be used as an ideal model to study the in vitro protein folding process. Accordingly, the effects of different conditions and molecules on the protein folding pathway can be efficiently studied using sGFP as a model protein.

Graphical abstract:

Schematic of the steps involved in the sGFP refolding pathway. Native sGFP is unfolded by chemical denaturation using 6 M GuHCl at 25°C for 1 hour and then refolded in refolding buffer by 100-fold dilution.

Small molecules that react to form covalent bonds with proteins are widely used as biological tools and therapeutic agents. Screening cysteine-reactive fragments against a protein target is an efficient way to identify chemical starting points for covalent probe development. Mass spectrometry is often used to identify the site and degree of covalent fragment binding. However, robust hit identification requires characterization of the kinetics of covalent binding that can be readily achieved using quantitative irreversible tethering. This screening platform uses a non-specific cysteine-reactive fluorogenic probe to monitor the rate of reaction between covalent fragments and cysteine containing biomolecules. Fragment libraries are simultaneously screened against the target protein and glutathione, which functions as a control, to identify hit fragments with kinetic selectivity for covalent modification of the target. Screening by quantitative irreversible tethering accounts for variations in the intrinsic reactivity of individual fragments enabling robust hit identification and ranking.