Mapping Protein-Protein Interactions
Mapping protein-protein interactions gives us a better understanding of molecular mechanisms inside the cell.
Compare techniques used for mapping protein-protein interactions
- A key question about a protein, in addition to when and where it is expressed, is with which other proteins does it interact?
- Interaction partners are an immediate lead into biological function and can potentially be exploited for therapeutic purposes.
- Creation of a protein–protein interaction map of the cell would be of immense value to understanding the biology of the cell.
- The most common approaches used to identify protein-protein interactions are the yeast two-hybrid system and affinity purification coupled to mass spectrometry.
- Affinity chromatography: Method of separating a biochemical mixture based on highly specific interactions.
- mass spectrometry: Analytical method that allows ionizing molecules and sorting them according to their mass and charge.
In living organisms most of the biological functions are mediated by complex multi-component protein machineries and network activities. The protein complexes formed could be stable (proteins interact for a prolonged period of time) or transient (proteins interact for a brief period of time). Molecular studies are necessary to dissect the constituents of these protein complexes and identify the domains through which a protein interacts with another. Understanding how proteins are physically connected reveals clues about their structure and function and makes them an ideal target for drug therapy. Several methodologies exist to study the interaction of proteins in vivo. The most widely employed tools are the yeast two-hybrid system and affinity purification coupled to mass spectrometry. Datasets obtained from such tools are further analyzed using computational methods to draw a map of protein connectivity and achieve system level understanding of a microorganism. The complete map of protein interactions that can occur in a living organism is called the interactome.
The Yeast Two-hybrid System
The yeast two-hybrid screening system is an effective and quick tool for the in vivo study of protein–protein interaction both in prokaryotes and eukaryotes. The method consists of splitting a yeast transcription factor into its binding domain and activation domain, fusing the binding domain to one protein of interest (the bait) and the activation domain to another protein of interest (the prey), and reconstituting the activity of the transcription factor by bringing the two domains back into physical proximity. In the absence of an interaction the domains remain distant, preventing a detectable output. If the two proteins do interact the bait recruits the prey to a specific cellular location where it can stimulate a detectable output (e.g., gene activation). This experimental approach measures direct physical interaction between proteins and is called a binary method.
Affinity Purification Coupled to Mass Spectrometry
Affinity purification of protein complexes coupled to mass spectrometry is carried out as follows: a specific protein (the bait) is manipulated to express an affinity tag. The tag serves as a tool to purify the bait protein and associated proteins by affinity chromatography. Purified protein complexes are then resolved on native gels and discrete protein bands are excised and digested into small peptide fragments by trypsin.
Peptides are identified using mass spectrometry methods. The identity of the protein associated with a given bait protein is determined by comparing its peptide fingerprint against available databases. This method allows for the identification and quantification of direct binding partners and secondary interacting proteins, and assigns them into protein networks. This experimental approach measures physical interactions between groups of proteins without distinguishing whether they are direct or indirect and is termed co-complex method. Results collected from binary and co-complex experiments are documented into a database. There are many databases accessible online that allow for protein clustering by function and nature of interaction and provide a rich framework for biomedical research.
Tracking Cells with Light
Advanced technology enables tracking cells with light by introducing fluorescent or luminescent reporter genes into the cells’ genome.
Compare the ways light can used to track cells
- The ability of tracking cells with light has revolutionized molecular biology and provided means to study biological processes as they happen.
- The most common tools used to illuminate a cell are fluorescent (GFP) and luminescent (luciferase) reporter genes.
- Reporter genes are introduced into the host ‘s genome and are controlled by the regulatory sequence of the gene under investigation (gene X). Thus when gene X is expressed it will drive along the expression of the reporter gene and the cell will fluoresce or emit light.
- Many laboratory devices are available to visualize illuminated living cells and these range from fluorescence microscopy to more advanced spectroscopy.
- Spectroscopy: use of light, sound or particle emission to study matter. The emissions provide information about the properties of the matter under investigation. The device often used for such analysis is a spectrometer, which records the spectrum of light emitted (or absorbed) by a given material.
- Fluorescence microscopy: optical microscope that uses fluorescence to study properties of substances. A sample is illuminated with light of a wavelength that excites fluorescence in the sample. The fluoresced light, which is usually at a longer wavelength than the illumination, is then imaged through a microscope objective.
Fluorescence and luminescence
Cells undergo many dynamic processes. In order to visualize these processes we need to be able to film cells over time. This can be achieved by using tools to monitor gene expression to track when proteins are made and where they go in the cell. In molecular biology, researchers use a reporter gene that they attach to a regulatory gene of interest. Reporter genes ideally have distinguishable properties that can be easily detected and measured. The most commonly used reporter genes have biofluorescent or bioluminescent characteristics and can be visualized with the aid of microscopy and other non-invasive imaging equipments. Examples of such reporters are the genes encoding for Green Fluorescent Protein (GFP) and luciferase, respectively. The discovery of GFP changed the way we look at cellular life today. GFP was first isolated from the jellyfish (Aequorea victoria) by the Japanese scientist Osuma Shimomura in the early 1960s. It was then cloned and its sequence identified in 1992 by Douglas Prasher. GFP is widely used in research laboratories as a marking tool to illuminate and track genes in fixed or living cells. Luciferase, isolated from fireflies, is an enzyme present in the cells of bioluminescent organisms that catalyzes the oxidation of luciferin and ATP producing light. Luciferase is similarly useful as a biological marker in living cells and organisms.
Transfection of reporter genes into cells
To introduce a reporter gene into an organism, scientists place the reporter gene and the gene of interest in the same DNA construct to be inserted into the cell or organism. For bacteria or prokaryotic cells in culture, this is usually in the form of a circular DNA molecule called a plasmid. It is important to use a reporter gene that is not natively expressed in the cell or organism under study, since the expression of the reporter is being used as a marker for successful uptake of the gene of interest. This gene’s regulatory sequence now controls the production of GFP or luciferase, in addition to the protein of interest. In cells where the gene is expressed, and the tagged proteins are produced, GFP or luciferase are produced at the same time. Thus, only those cells in which the tagged gene is expressed, or the target proteins are produced, will fluoresce when observed under fluorescence microscopy, or bioluminesce (emit light) when luciferin, the substrate for luciferase is added.
Application of GFP in molecular microbiology
GFP has many advantages over conventional reporter genes in that it is highly stable, non-toxic to living cells and organisms, detection tools are non-invasive and the green light is generated without the addition of external cofactors and measured without application of expensive equipment. Various applications of that reporter gene were documented and vary from being able to monitor microorganism ‘s survival in complex biological systems such as activated sludge to biodegradation of chemical compounds in soil. GFP allowed the detection, determination of spatial location and enumeration of bacterial cells from diverse environmental samples such as biofilm and water. GFP as biomarker is also useful in monitoring gene expression and protein localisation in bacterial cells.
Multiplex and Real-Time PCR
Multiplex and real-time PCR are molecular techniques designed to amplify nucleic acid sequences in a quantitative manner.
Illustrate the use and method of multiplex and real-time PCR
- Real-time PCR is a molecular tool for nucleic acid amplification monitored as the reaction progresses.
- Multiplex PCR technique can use fluorescence to detect, quantitate, and visualize PCR products on a computer monitor by utilizing numerous primer sets.
- Real-time PCR can be a simplex, amplifying one DNA template with one set of primers, or multiplex, amplifying one or more DNA templates with one or more sets of primers in one reaction.
- agarose gel electrophoresis: Method used for the separation of DNA fragments by size.
- oligonucleotide: A strand of nucleic acid that serves as a starting point for DNA synthesis.
Polymerase Chain Reaction (PCR) is a molecular technique commonly used to amplify nucleic acid sequences. The starting material is a messenger RNA (mRNA) of interest that could be obtained from a wide array of sample types and extracted using commercially available kits and reagents. This mRNA is used to synthesize complementary DNA (cDNA) in a reaction catalyzed by the enzyme reverse transcriptase. The importance of this step is it allows converting a labile RNA into its more stable cDNA form that can be stored and used for multiple applications. The resulting cDNA serves as the template for the PCR reaction. The PCR process can be divided into three steps: DNA denaturation where double-stranded DNA (dsDNA) is separated at temperatures above 90°C, oligonucleotide primers annealing at 50–60°C, and primer extension at 70–78°C. A programmable thermal cycler controls the rate of temperature change, the length of the incubation at each temperature, and the number of times each cycle of temperatures is repeated. The final product of the reaction is called amplicon. It is confirmed by agarose gel electrophoresis for qualitative results.
Real-time polymerase chain (RT-PCR) reaction, also called quantitative real-time PCR (qRt-PCR) is used to amplify and quantify targeted DNA molecules. The use of RT-PCR allows for both detection and quantitation of DNA sequences. The quantity can be an absolute number of copies or a relative amount when normalized to DNA input or additional normalizing genes. The procedure for RT-PCR follows the general principles of PCR, but the defining feature is the ability to detect amplified DNA as the reaction progresses in real time.
Real-time PCR can used to amplify low-abundance DNA templates. It is useful in monitoring the accumulating amplicon. Two common methods that are used to product detection in real-time PCR include the use of non-specific flourescent dyes that intercalate with double-stranded DNA or sequence-specific DNA probes that consist of oligonucleotides labeled with a fluorescent reporter (oligoprobes). The fluorescent reporter permits detection after hybridization of the probe with its complementary DNA target. During real-time PCR with oligoprobes, there is a change in signal following direct interaction with the amplicon. The signal is related to the amount of amplicon present during each cycle and will increase as the amount of specific amplicon increases. The detection of amplicon could be visualized on a graph as the amplification progresses. Real-time PCR assays have been extremely useful for studying microbial agents of infectious disease and have proven valuable for basic microbiological research. The ability to amplify templates from a broad selection of specimen has made it an ideal system for application across the various microbiological disciplines.
A new and improved technology called multiplex PCR was introduced to allow the use of one or more primer sets to potentially amplify multiple templates within a single reaction. Up to 20 different reactions can be run simultaneously, therefore lowering the amount of sample used, reducing the reagents consumed, and collecting far more information per reaction, while simplifying data analyses. Multiplex PCR is a challenging application that typically requires more optimization than standard, single amplicon PCR assays. The key to successful multiplex PCR is the ability to define a single set of reaction parameters (reagent concentrations and cycling parameters) that allows for all primers to anneal with high specificity to their target sequences and be extended with the same efficiency. Primer design, as well as the enzyme and buffer system, are critical factors in this challenge. The results from multiplex PCR can be analyzed using gel electrophoresis or using fluorophores for analysis using during the reaction. Ideally, a real-time multiplex PCR should be able to detect, differentiate, and provide a quantitative result for many different targets without a single target influencing the detection of one of the others (cross-talk) and without loss of sensitivity. It is evident that due to the limited number of fluorophoric labels available and the significant overlap in their emission spectra, quantification of multiplex reaction products is often difficult.
Numerous companies have helped overcome this issue by making dyes available that are compatible for use in multiplex PCR. Since its first description in 1988 by Chamberlain et al, this method has been applied in many areas of DNA testing, including analyses of deletions, mutations, and polymorphisms, or quantitative assays and reverse transcription PCR. Typically, it is used for genotyping applications where simultaneous analysis of multiple markers is required, detection of pathogens or genetically modified organisms, or for microsatellite analyses. Multiplex assays can be tedious and time-consuming to establish, requiring lengthy optimization procedures but once optimized numerous high-throughput genomic assays can be achieved at optimum speed.
Phage display is a laboratory tool based on cloning DNA sequences into a phage which presents proteins encoded by that DNA on its surface.
Assess the uses of phage display technology
- A phage, short for bacteriophage, is a virus that reproduces itself in bacteria.
- Phage display technology introduces genes into the phage’s genome which encoded proteins would be presented on the surface of the phage.
- Displayed proteins are tested for binding affinity against target molecules immobilized on a platform.
- lysozyme: enzyme that damages bacterial cell wall.
- sequencing: the process of reading the nucleotide bases in a DNA molecule.
Definition of a bacteriophage
A phage or bacteriophage is a virus capable of infecting a bacterial cell, and may cause lysis to its host cell. Bacteriophages have a specific affinity for bacteria. They are made of an outer protein coat or capsid that encloses the genetic material (which can be RNA or DNA, about 5,000 to 500,000 nucleotides in length). They inject their genetic material into the bacterium following infection. When the strain is virulent, all the synthesis of the host’s DNA, RNA and proteins ceases. The phage genome is then used to direct the synthesis of phage nucleic acids and proteins using the host’s transcriptional and translational apparatus. When the sub-components of the phage are produced, they self-assemble to form new phage particles. The new phages produce lysozyme that ruptures the cell wall of the host, leading to the release of the new phages, each ready to invade other bacterial cells. This inherent property of phages is the basis for the phage display technology.
Phage display technology
Phage display technology is the process of inserting new genetic material into a phage gene. The bacteria process the new gene so that a new protein or peptide is made. This protein or peptide is exposed on the phage surface. Phage display begins by inserting a diverse set of genes into the phage genome with each phage receiving a different gene. The modified gene contains an added segment (an antibody, small protein, or peptide), which is to be expressed on the surface of the phage. Each phage receives only one gene, so each expresses a single protein or peptide. A collection of phage displaying a population of related but diverse proteins or peptides is called a library. The related proteins keep most of the physical and chemical properties of their parent protein. The library is then exposed to an immobilized target. It is anticipated that some members of the library will bind to the target through an interaction between the displayed molecule and the target itself. After the phage is given the chance to bind to a target, the immobilized target is washed to remove phage that did not bind. Replicating the bound phage in bacteria increases the amount of phage several million-fold overnight, providing enough material for sequencing. Sequencing of the phage DNA tells the identity of the peptide that binds the target. Phage libraries are screened for binding to synthetic or native targets.
Phage display technology is advantageous in many applications including selection of inhibitors for the active and allosteric sites of enzymes, receptor agonists and antagonists, and G-protein binding modulatory peptides. Phage display is also used in epitope mapping and analysis of protein-protein interactions. The specific molecules isolated from phage libraries can be used in therapeutic target validation, drug design and vaccine development.