Proteins in the cell are in a constant flux governed by events including synthesis and degradation. In an effort to make cells more efficient by reducing the unnecessary protein load, most proteins in the cell have a specifically defined half-life. Another reason why cells have evolved to degrade proteins is to ensure timely removal of proteins that have accomplished their job (cyclins for example) and prevent undesirable effects that may occur due to the constitutive action of some proteins (for instance, deregulation in proteasomal degradation of cell cycle proteins fueling uncontrolled cancerous proliferation).
Have you ever wondered how stable your protein is intracellularly? If your mentor or graduate committee was to ask you what is the intracellular half-life of your protein, then what experiments would you need to do to come up with to answer that? Here, I will introduce you some common methods that are employed to measure protein stability.
Traditionally, this was the method of choice to track proteins and estimate their stability. Barring the hassle that involves working with radioactivity, this method holds a distinct advantage over other pharmacological ways of measuring protein half-life and is still frequently used in the lab. Not only does it allow you to measure protein half-life but if you are really skilled at autoradiography and subcellular fractionation, you could also track the protein and determine its subcellular localization. The experiment involves labeling cells with radioactive precursors (pulse, typically done by adding radioactive methionine and/or cysteine to the media) for a short duration and then following the protein kinetics as cells are allowed to recover in media supplemented with unlabeled precursors (called excess cold precursor). As the old protein bearing the labeled precursor is getting degraded within the cells, the new protein is concurrently being synthesized using the unlabeled precursors. The protein is then immunoprecipitated at different time points and radioactivity in the sample is measured as a function of time (chase) by autoradiography. Based on how fast the non-radioactive protein replaces the radioactive one, one can determine the half-life of any protein. Another non-radioactive variation of this method that has become fashionable recently is the bleach-chase method. In this method, the protein of interest is fused with fluorophore and the protein population is bleached with a brief pulse of light, producing two subsets of protein population – fluorescently and non-fluorescently population. The rate of fluorescence recovery post-bleaching is then correlated to rate of degradation.
This provides a much easier alternative to the pulse-chase and avoids you having to work with radioactivity. Typically, the method involves arresting protein synthesis by adding Cycloheximide and then assessing the degradation profile of your favorite protein (YFP) as a function of time. Cycloheximide is de-novo protein synthesis inhibitor that acts by inhibit translation elongation through binding to the E-site of the 60S ribosomal unit. This, in turn, arrests protein synthesis allowing investigators to follow degradation profile of proteins without having to worry about the effects of ongoing protein synthesis. Think of it as it as a long conveyer belt in any factory. If you want to know how fast unloading is occurring (degradation), you can’t have workers simultaneously loading it with products at the manufacturing end (synthesis). When you stop the loading of new products on this belt, the unloading speed can be assessed by finding out how long the belt runs (time), and how many packets (protein) are unloaded during that time. When you divide the number of packets unloaded by the amount of time the belt ran, this will give you the unloading speed in packets per unit time (degradation rate). In other words, adding Cyclohexmide takes out the protein synthesis components out of the equation allowing investigators to exclusively study degradation of proteins as a function of time.
Pharmacological Inhibitors of Degradation Pathways
Once you have determined the rate of your degradation you will want to know the mechanism. There are two main degradation pathways inside the cell: The proteasomal pathway and the lysosomal pathway. There are specific pharmacological inhibitors available for assessing degradation by either route. The proteasomal pathway involves tagging the protein with ubiquitin by a series of enzymes. This ubiquitin-tagged protein is then destined towards the cellular proteasomal complex where it is degraded. However, proceed with caution because in some cases proteasomal degradation may occur independently of ubiquitination and this is a scenario one must not disregard when testing a hypothesis regarding possible degradation routes. To make matters even more complicated, ubiquitination may also play a role in the lysosomal route of degradation. Lysosomal degradation is also sub-classified into many different types based on subtleties in actual cellular mechanisms of degradation of the cargo (macroautophagy, microautophagy and chaperone-mediated autophagy). To assess if your protein is getting degraded through the proteasomal pathway you can pharmacologically inhibit the proteasomal complex and assess the effect of this intervention on YFP expression. Investigators typically use proteasomal inhibitors like MG-132, Epoxomicin, and Bortezomib. These inhibitors bind to specific proteolytic sites within the proteasomal complex and prevent the degradation of proteins. There is also an arsenal of pharmacological inhibitors and genetic strategies to study autophagic degradation, a topic that merits a separate discussion. Examples of some inhibitors used to study this mode of degradation include drugs like Chloroquine and BafimlmycinA1 that act by neutralizing the acidic pH in lysosomes required for action of lysosomal proteases. You would want to establish the relevant dose and duration of treatment and concurrently check the viability of your cells to ensure that you don’t kill the cells if you treat the cells with these drugs.
Mining Degradation Relevant Post-translational Modifications
Once you determine the degradation route and pathway responsible for YFP, you could analyze which tag (post-translational modification, PTM) is necessary for this process. There are many types of PTMs that can tag the proteins for degradation. There are also specific degradation motifs (affectionately called degrons) that target proteins towards degradation pathway. Some common examples of these degrons include D-boxes, PEST, and KEN boxes. Such degrons can assist the tagging of protein and facilitate their degradation through various mechanisms. For instance some of these degrons may make the protein more accessible to E3 ligase which can then escort it to the proteasomal complex.
One can check the primary amino acid sequence of the protein to identify if there are any specific degrons that jump out at you. If there is no precedence in the literature for degradation relevant PTMs there are specific databases one can quickly check before embarking on a mutational analysis of your protein. Such databases house sequences identified from large high-throughput proteomic studies where investigators have cataloged PTMs derived from global degradation profiles. Such analysis could provide you with crucial hints regarding degradation relevant PTM for YFP, allowing you to narrow down the region within a protein necessary for degradation, and if you are lucky to even map out the residue that plays a role in degradation. Examples of such databases include PhosphoSite (phosphorylation and other PTMs) and mUbiSiDa (ubiquitination).
After narrowing down the residues you think are relevant for degradation, the next stage is changing these residues and assessing the effect of such mutations on the stability of YFP. For instance, a lysine to arginine mutation on a degradation relevant site can get rid of the epsilon amino group that is necessary for ubiquitin attachment and can stabilize the protein. Similarly, if you believe phosphorylation is a crucial event for degradation, phospho-defective (Serine to Alanine for instance) and phospho-mimicking mutations (serine to glutamic acid for instance) can alter the degradation profile of YFP.
Analyzing Ubiquitination Kinetics of Your Protein
Proteasomal degradation is often preceded by ubiquitination. Detecting ubiquitination can provide a strong piece of data corroborating that your protein is most likely getting targeted to the proteasome. These experiments involve pulling down YFP and then assessing its ubiquitination status using anti-ubiquitin antibody. What you would typically expect to see is a smear or a ladder-like banding effect when your IP samples are probed with anti-ubiquitin antibody. This happens because different higher-modified forms of the proteins are recognized as they progressively get tagged with multiple ubiquitin molecules. The intensity of the detected ubiquitin smears may get darker following proteasomal block – indicating an accumulation in polyubiquitinated forms of your protein. Ubiquitinated proteins can be notoriously unstable and specific experimental conditions and time of harvesting the cells must be carefully optimized to prevent their loss during sample prep. Since in many cases the read-out in this assay is a smear, an antibody-based method may produce artifacts due to non-specific pulldowns. A better and cleaner approach is tagging your protein of interest with hexahistidine tag (his-tag) and using Ni-affinity pulldown to assess ubiquitination on the protein. Another good control is using an HA-tagged version of ubiquitin plasmids and co-transfecting it with your protein of interest prior to pulldown. Eliminating one or both of these plasmids should decrease or even get rid of ubiquitin smears providing confidence in your ubiquitination assay.
These assays provide you with a cool tool-kit to study protein degradation and produce that awesome biochemistry paper you always wanted!