Biochemistry

Proteolysis is a critical biochemical process yet a challenging field to study experimentally due to the self-degradation of a protease and the complex, dynamic degradation steps of a substrate. Mass spectrometry (MS) is the traditional way for proteolytic studies, yet it is challenging when time-resolved, step-by-step details of the degradation process are needed. We recently found a way to resolve the cleavage site, preference/selectivity of cleavage regions, and proteolytic kinetics by combining site-directed spin labeling (SDSL) of protein substrate, time-resolved two-dimensional (2D) electron paramagnetic resonance (EPR) spectroscopy, protease immobilization via metal–organic materials (MOMs), and MS. The method has been demonstrated on a model substrate and protease, yet there is a lack of details on the practical operations to carry out our strategy. Thus, this protocol summarizes the key steps and considerations when carrying out the EPR/MS study on proteolytic processes, which can be generalized to study other protein/polypeptide substrates in proteolysis. Details for the experimental operation and cautions of each step are reported with figures illustrating the concepts. This protocol provides an effective approach to understanding the proteolytic process with the advantages of offering time-resolved, residue-level resolution of structural basis underlying the process. Such information is important for revealing the cleavage site and proteolytic mechanisms of unknown proteases. The advantage of EPR, probing the target substrate regardless of the complexities caused by the proteases and their self-degradation, offers a practically effective, rapid, and easy-to-operate approach to studying proteolysis.

Key features

• Combining protease immobilization, EPR, spin labeling, and MS experimental methods allows for the analysis of proteolysis process in real time.

• Reveals cleavage site, kinetics of product generation, and preference of cleavage regions via time-resolved SDSL-EPR.

• MS confirms EPR findings and helps depict the sequences and populations of the cleaved segments in real time.

• The demonstrated method can be generalized to other proteins or polypeptide substrates upon proteolysis by other proteases.

Graphical overview

Eukaryotic cells have different types of proteasomes that differ in size. The smallest proteolytically active particle is the 20S proteasome, which degrades damaged and oxidized proteins; the most common larger particle is the 26S proteasome, which degrades ubiquitylated proteins. The 26S proteasome is formed by a 20S particle capped with one or two regulatory particles, named 19S. While proteasome particles function in the cytoplasm, endoplasmic reticulum, and nucleus, our understanding of their abundance and activity in different cellular compartments is still limited. We provide a three-step protocol that first involves detergent-based fractionation of the cytoplasmic and nuclear compartments, maintaining the integrity and activity of proteasome complexes. Second, the protocol employs native gel separation of large multiprotein complexes in the fractions and a fluorescence-based in-gel quantitation of the activity and different proteasome particles. Finally, the protocol involves protein in-gel denaturation and transfer to a PVDF membrane. Western blotting then detects and quantifies the different proteasome particles. Therefore, the protocol allows for sensitive measurements of activity and abundance of individual proteasome particles from different cellular compartments. It has been optimized for motor neurons induced from mouse embryonic stem cells but can be applied to a variety of mammalian cell lines.

Key features

• Protocol for fractionation of active nuclear and cytoplasmic proteasome complexes.

• Native electrophoresis and fluorescence-based in-gel activity assay, which allows the visualization and quantification of active complexes within the acrylamide gel matrix.

• In-gel protein denaturation followed by transfer of complexes to PVDF membrane, which allows the analysis of complexes’ abundance using antibodies.

Graphical overview

Cycloheximide (CHX) is a small molecule derived from Streptomyces griseus that acts as fungicide. As a ribosome inhibitor, CHX can restrict the translation elongation of eukaryotic protein synthesis. Once protein synthesis is inhibited by CHX, the level of intracellular proteins decreases by degradation through the proteasome or lysosome system. Thus, the CHX chase assay is widely recognized and used to observe intracellular protein degradation and to determine the half-life of a given protein in eukaryotes. Here, we present a complete experimental procedure of the CHX chase assay.

Graphical overview

In this protocol, we describe the analysis of protein stability over time, using synthesis shutoff. As an example, we express HA-tagged yeast mitofusin Fzo1 in Saccharomyces cerevisiae and inhibit translation via cycloheximide (CHX). Proteasomal inhibition with MG132 is performed, as an optional step, before the addition of CHX. Proteins are extracted via trichloroacetic acid (TCA) precipitation and subsequently separated via SDS-PAGE. Immunoblotting and antibody-decoration are performed to detect Fzo1 using HA-specific antibodies. We have adapted the method of blocking protein translation with cycloheximide to analyze the stability of high molecular weight proteins, including post-translational modifications and their impact on protein turnover.

Fibrinolysis is an integral part of the matrix remodeling process that contributes to tissue repair. Fibrin clots are broken down during fibrinolysis in a controlled process. Fibrin degradation products (FDPs) have also been shown to have a role in the regulation of cell growth and are implicated in various vascular diseases. This protocol was designed to quantitatively measure the extent of fibrin breakdown and how this can be adapted as a tool to further investigate the pathway involved in fibrinolysis or fibrin degradation products. Until now, we haven’t found an alternative method to analysis fibrinolysis.

Cyclic nucleotide degrading phosphodiesterase (PDE) enzymes are crucial to the fine tuning of cAMP signaling responses, playing a pivotal role in regulating the temporal and spatial characteristics of discrete cAMP nanodomains and hence the activity of cAMP-effector proteins. As a consequence of orchestrating cAMP homeostasis, dysfunctional PDE activity plays a central role in disease pathogenesis. This highlights the need for developing methods that can be used to further understand PDE function and assess the effectiveness of potentially novel PDE therapeutics. Here we describe such an approach, where PDE activity is indirectly measured through the direct quantification of radioactively tagged cAMP (pmol/min^-1/mg^-1). This method provides a highly sensitive tool for investigating PDE functionality.

Autophagy is a key player in the maintenance of cellular homeostasis in eukaryotes, and numerous diseases, including cancer and neurodegenerative disorders, are associated with alterations in autophagy. The interest for studying autophagy has grown intensely in the last two decades, and so has the arsenal of methods utilised to study this highly dynamic and complex process. Changes in the expression and/or localisation of autophagy-related proteins are frequently assessed by Western blot and various microscopy techniques. Such analyses may be indicative of alterations in autophagy-related processes and informative about the specific marker being investigated. However, since these proteins are part of the autophagic machinery, and not autophagic cargo, they cannot be used to draw conclusions regarding autophagic cargo flux. Here, we provide a protocol to quantitatively assess bulk autophagic flux by employing the long-lived protein degradation assay. Our procedure, which traces the degradation of ¹⁴C valine-labelled proteins, is simple and quick, allows for processing of a relatively large number of samples in parallel, and can in principle be used with any adherent cell line. Most importantly, it enables quantitative measurements of endogenous cargo flux through the autophagic pathway. As such, it is one of the gold standards for studying autophagic activity.

The fine-tuned balance of protein level, conformation and location within the cell is vital for the dynamic changes required for a cell to respond to a given stimulus. This requires the regulated turnover of damaged or short-lived proteins through the ubiquitin proteasome system (UPS). Thus, the protease activity of the proteasome is adjusted to meet the current demands of protein degradation via the UPS within the cell. We describe the adaptation of an intramolecular quenched fluorescence assay utilizing substrate-mimic peptides for the measurement of proteasome activity in total plant extracts. The peptide substrates contain donor-quencher pairs that flank the scissile bond. Following cleavage, the increase in dequenched donor emission of the product is subsequently measured over time and used to calculate the relative proteasome activity.

Highly regulated and targeted protein degradation plays a fundamental role in almost all cellular processes. Determination of the protein half-life by the chase assay serves as a powerful and popular strategy to compare the protein stability and study proteolysis pathways in cells. Here, we describe a chase assay in Haloferax volcanii, a halophilic archaeon as the model organism.

The half-life of a protein is a characteristic property, and can be modulated by post-translational modifications, changes in subcellular localization, and/or interaction with other proteins or ligands. As one determinant of its steady-state level, a protein’s degradation represents an important distinguishing attribute relevant to its biological function. Because protein longevity cannot be elucidated from bioinformatics analyses, it must be determined empirically. Here we describe two approaches for in vivo half-life determination in plants: 1. pooled-seedling degradation assays monitoring either tagged versions of the protein (luciferase fusions or other epitope tags) or following the endogenous protein; 2. single-seedling degradation assays using luciferase fusion proteins. The advantages of these approaches are their simplicity and low cost.