Studying protein function in live cells is limited by the tools available to analyze the expression and interactions of those proteins. Although mass spectrometry and antibody-based protein detection are valuable technologies for protein analysis, both methods have drawbacks that limit the range of targets and contexts in which proteins can be investigated.
Mass spectrometry is often poor at detecting low-abundance proteins. Antibody-based techniques require high quality, specific antibodies, which can be difficult to impossible to acquire. Both methods require cell lysis, preventing real-time analysis and limiting the physiological relevance, and both methods can be limiting for higher-throughput analysis. While plasmid-based overexpression of tagged target proteins simplifies detection and can allow for real time analysis, protein levels don’t typically resemble endogenous levels. Overexpression also has the potential to create experimental artifacts or limit the dynamic range of an observed response.
While their findings showed that this method provides efficient and specific tagging of endogenous proteins, the research was limited to just five different proteins within a single signaling pathway in two cell lines. This left unanswered questions about whether this approach was scalable, had broader applications and how accurately the natural biology of the cells was represented.
Transcriptional activation of genes within the nucleus of eukaryotic cells occurs by a variety of mechanisms. Typically, these mechanisms rely on the interaction of regulatory proteins (transcriptional activators or repressors) with specific DNA sequences that control gene expression. Upon DNA binding, regulatory proteins also interact with other proteins that are part of the RNA polymerase II transcriptional complex.
One type of transcriptional activation relies on inducing a conformational change in chromatin, the DNA-protein complex that makes up each chromosome within a cell. In a broad sense, “extended” or loosely wound chromatin is more accessible to transcription factors and can signify an actively transcribed gene. In contrast, “condensed” chromatin hinders access to transcription factors and is characteristic of a transcriptionally inactive state. Acetylation of lysine residues in histones—the primary constituents of the chromatin backbone—results in opening up the chromatin and consequent gene activation. Disruption of histone acetylation pathways is implicated in many types of cancer (1).
Recently, Gordon et al. published an atlas of protein:protein interactions of all proposed SARS-CoV-2 proteins expressed individually in HEK 293 cells (Table 1). The study tagged each of the viral proteins with an epitope tag and performed a pull-down of the expressed protein followed by trypsin digestion and mass spec analysis, a process referred to as affinity purification–mass spec analysis. The group identified 332 human proteins interacting with 27 SARS-CoV-2 proteins.
The interactions identified in the HEK 293 cells helped Appelberg et al. analyze interactions over time in SARS-CoV-2-infected Huh7 cells. Gordon et al. used the PPI data to identify FDA-approved drugs, drugs in clinical trials, and pre-clinical compounds that bound to the identified human proteins and labs in New York and Paris tested some of these drugs for antiviral effects.
NanoLuc® luciferase has been discussed many times on this blog and our web site because the enzyme is integral to studying genetic responses and protein dynamics. While NanoLuc® luciferase was first introduced as a reporter enzyme to assess promoter activity, its capabilities have expanded far beyond a genetic reporter, creating tools used to study endogeneous protein interactions, target engagement, protein degradation and more. So where did the NanoLuc® luciferase come from and how does a one enzyme power several technologies?
The ability to target protein interactions with low solubility or weak binding affinities can present a significant challenge when it comes to drug screening. The beauty of these types of challenges we often face in the lab is that finding solutions to these problems doesn’t necessarily require brand new tools. Sometimes we already have the right tools in our arsenal and, with just a little creativity and collaboration, they can be adapted to address the challenge at hand.
In the following video, Dr. Mohamed (Soly) Ismail, a Postdoctoral Fellow at the Downward Lab of the Francis Crick Institute, presents the perfect example of this with his novel approach to the NanoBiT® Protein:Protein Interaction Assay. Through a collaboration with Promega R&D Scientists, Dr. Ismail has translated the assay into a cell-free, biochemical format, termed the NanoBiT Biochemical Assay (NBBA).
Do you find the thought of a giant rodent off-putting? Do your thoughts go to huge rats running amuck in dark allies, threatening unsuspecting passers by?
I personally hold rodents in low esteem. Rats, mice…who needs them? With the exception of cavies. I spent countless hours as a child playing with guinea pigs. We had as many as 16 of these little rodents at one time (the males are very capable of chewing or climbing out of cardboard boxes to reach a female in the next box). The baby guinea pigs were very cute and the adults had quite pronounced personalities, and a lot of attitude.
It was this history with guinea pigs that made me interested in learning more about the largest rodent in the world, the South American capybara (Hydrochoerus hydrochaeris). These family-oriented herbivores are found in savannas and forested areas, living in groups of as many as 100 members. They are excellent swimmers and can remain underwater for as long as 5 minutes. In fact, capybara mate only in the water. (Perhaps it’s not surprising then that the South American alligator, the caiman, is one of the capybara’s greatest predators.)
With their squared-off nose and lack of tail, capybaras actually resemble guinea pigs. However, these oversized cavies weigh as much as 40 pounds. and can reach 24” at the shoulder, the size of an average standard poodle. Guinea pigs, on the other hand, weigh in at 2–3 pounds, and are 3–4” tall.
Their proportions make capybaras 60 times more massive than their closest relatives, rock cavies (Kerodon sp.) and 2,000 times more massive than the common mouse (Mus musculus). This tremendous size difference is why Herrera-Álvarez et al. took a closer look at the capybara, studying its propensity to develop cancer and other tradeoffs that would seem to coincide with its exceptional size.
Our cells have evolved multiple mechanisms for “taking out
the trash”—breaking down and disposing of cellular components that are defective,
damaged or no longer required. Within a cell, these processes are balanced by
the synthesis of new components, so that DNA, RNA and proteins are constantly
Proteins are degraded by two major components of the cellular
machinery. The discovery of the lysosome in the mid-1950s
provided considerable insight into the first of these degradation mechanisms
for extracellular and cytosolic proteins. Over the next several decades,
details of a second protein degradation mechanism emerged: the ubiquitin-proteasome system
(UPS). Ubiquitin is a small, highly conserved polypeptide that is used to
selectively tag proteins for degradation within the cell. Multiple ubiquitin
tags are generally attached to a single targeted protein. This ill-fated, ubiquitinated
protein is then recognized by the proteasome, a large protein complex with
proteolytic activity. Ubiquitination is a multistep process, involving several
specialized enzymes. The final step in the process is mediated by a family of ubiquitin
ligases, known as E3.
Understanding the expression, function and dynamics of
proteins in their native environment is a fundamental goal that’s common to
diverse aspects of molecular and cell biology. To study a protein, it must
first be labeled—either directly or indirectly—with a “tag” that allows
specific and sensitive detection.
Using a labeled antibody to the protein of interest is a
common method to study native proteins. However, antibody-based assays, such as
ELISAs and Western blots, are not suitable for use in live cells. These
techniques are also limited by throughput and sensitivity. Further, suitable
antibodies may not be available for the target protein of interest.
Synthetic biology—genetically engineering an organism to do or make something useful—is the central goal of the iGEM competition each year. After teams conquer the challenge of cloning their gene, the next hurdle is demonstrating that the engineered gene is expressing the desired protein (and possibly quantifying the level of expression), which they may do using a reporter gene.
Reporters can also play a more significant role in iGEM projects when teams design their organism with reporter genes to detect and signal the presence of specific molecules, like environmental toxins or biomarkers. Three of the iGEM teams Promega sponsored this year opted to incorporate some version of NanoLuc® Luciferase into their projects.
NanoLuc® luciferase is a small monomeric enzyme (19.1kDa, 171 amino acids) based on the luciferase from the deep sea shrimp Oplophorus gracilirostris. This engineered enzyme uses a novel substrate, furimazine, to produce high-intensity, glow-type luminescence in an ATP-independent reaction. Unlike other molecules for tagging and detecting proteins, NanoLuc® luciferase is less likely to interfere with enzyme activity and affect protein production due to its small size.
NanoLuc® Luciferase has also been engineered into a structural complementation reporter system, NanoBiT® Luciferase, that contains a Large subunit (LgBiT) and two small subunit options: low affinity SmBiT and high affinity HiBiT. Together, these NanoLuc® technologies provide a bioluminescent toolbox that was used by the iGEM teams to address a diverse set of biological challenges.
Here is an overview of each team’s project and how they
incorporated NanoLuc® technology.
You have identified and cloned your protein of interest, but you want to explore its function. A protein fusion tag might help with your investigation. However, choosing a tag for your protein depends on what experiments you are planning. Do you want to purify the protein? Would you like to identify interacting proteins by performing pull-down assays? Are you interested in examining the endogenous biology of the protein? Here we cover the advantages and disadvantages of some protein tags to help you select the one that best suits your needs.
The most commonly used protein tags fall under the category of affinity tags. This means that the tag binds to another molecule or metal ion, making it easy to purify or pull down your protein of interest. In all cases, the tag will be fused to your protein of interest at either the amino (N) or carboxy (C) terminus by cloning into an expression vector. This protein fusion can then be expressed in cells or cell-free systems, depending on the promoter the vector contains. Continue reading “Choosing a Tag for Your Protein”