3.1 Protein Purification3.2 Protein Identification and Visualization3.3 Protein Synthesis and Sequencing3.4 Protein Structure Elucidation3.5 Proteome Analysis3.6 References3.1 Protein PurificationProtein purification is a series of processes intended to isolate one or a few proteins from a complex mixture, usually cells, tissues or whole organisms. Protein purification is vital for the characterization of the function, structure and interactions of the protein of interest. The purification process may separate the protein and non-protein parts of the mixture, and finally separate the desired protein from all other proteins. Separation of one protein from all others is typically the most laborious aspect of protein purification. Separation steps usually exploit differences in protein size, physico-chemical properties, binding affinity and biological activity. The pure result may be termed protein isolate. Show
Protein purification is either preparative or analytical. Preparative purifications aim to produce a relatively large quantity of purified proteins for subsequent use. Examples include the preparation of commercial products such as enzymes (e.g. lactase), nutritional proteins (e.g. soy protein isolate), and certain biopharmaceuticals (e.g. insulin). Several preparative purifications steps are often deployed to remove bi-products, such as host cell proteins, which poses as a potential threat to the patient’s health. Analytical purification produces a relatively small amount of a protein for a variety of research or analytical purposes, including identification, quantification, and studies of the protein’s structure, post-translational modifications and function. Pepsin and urease were the first proteins purified to the point that they could be crystallized. ExtractionIf the protein of interest is not secreted by the organism into the surrounding solution, the first step of each purification process is the disruption of the cells containing the protein. Depending on how fragile the protein is and how stable the cells are, one could, for instance, use one of the following methods: i) repeated freezing and thawing, ii) sonication, iii) homogenization by high pressure (French press), iv) homogenization by grinding (bead mill), and v) permeabilization by detergents (e.g. Triton X-100) and/or enzymes (e.g. lysozyme). Finally, the cell debris can be removed by centrifugation so that the proteins and other soluble compounds remain in the supernatant. Also proteases are released during cell lysis, which will start digesting the proteins in the solution. If the protein of interest is sensitive to proteolysis, it is recommended to proceed quickly, and to keep the extract cooled, to slow down the digestion. Alternatively, one or more protease inhibitors can be added to the lysis buffer immediately before cell disruption. Sometimes it is also necessary to add DNAse in order to reduce the viscosity of the cell lysate caused by a high DNA content. Precipitation and Differential SolubilizationIn bulk protein purification, a common first step to isolate proteins is precipitation using a salt such as ammonium sulfate (NH4)2SO4. This process is called Salting In or Salting Out (Figure 3.1) This is performed by adding increasing amounts of ammonium sulfate and collecting the different fractions of precipitate protein. Ammonium sulfate is often used as it is highly soluble in water, has relative freedom from temperature effects and typically is not harmful to most proteins. Furthermore, ammonium sulfate can be removed by dialysis (Figure 3.2). The hydrophobic groups on the proteins get exposed to the atmosphere, attract other protein hydrophobic groups and get aggregated. Protein precipitated will be large enough to be visible. One advantage of this method is that it can be performed inexpensively with very large volumes. Figure 3.1 Salting In and Salting Out. During the salting in process, salt molecules increase the solubility of proteins by reducing the electrostatic interactions between protein molecules. As the salt concentration is increased, protein-protein interactions become more energetically favorable than protein-solvent interactions and the proteins precipitate from solution. Image derived from Michel Awkal The first proteins to be purified are water-soluble proteins. Purification of integral membrane proteins requires disruption of the cell membrane in order to isolate any one particular protein from others that are in the same membrane compartment. Sometimes a particular membrane fraction can be isolated first, such as isolating mitochondria from cells before purifying a protein located in a mitochondrial membrane. A detergent such as sodium dodecyl sulfate (SDS) can be used to dissolve cell membranes and keep membrane proteins in solution during purification; however, because SDS causes denaturation, milder detergents such as Triton X-100 or CHAPS can be used to retain the protein’s native conformation during complete purification. Figure 3.2 Dialysis. The process of dialysis separates dissolved molecules by their size. The biological sample is placed inside a closed membrane, where the protein of interest is too large to pass through the pores of the membrane, but through which smaller ions can easily pass. As the solution comes to equilibrium, the ions become evenly distributed throughout the entire solution, while the protein remains concentrated in the membrane. This reduces the overall salt concentration of the suspension. Image adapted from Gjk003 UltracentrifugationCentrifugation is a process that uses centrifugal force to separate mixtures of particles of varying masses or densities suspended in a liquid. When a vessel (typically a tube or bottle) containing a mixture of proteins or other particulate matter, such as bacterial cells, is rotated at high speeds, the inertia of each particle yields a force in the direction of the particles velocity that is proportional to its mass. The tendency of a given particle to move through the liquid because of this force is offset by the resistance the liquid exerts on the particle. The net effect of “spinning” the sample in a centrifuge is that massive, small, and dense particles move outward faster than less massive particles or particles with more “drag” in the liquid. When suspensions of particles are “spun” in a centrifuge, a “pellet” may form at the bottom of the vessel that is enriched for the most massive particles with low drag in the liquid. Non-compacted particles remain mostly in the liquid called “supernatant” and can be removed from the vessel thereby separating the supernatant from the pellet. The rate of centrifugation is determined by the angular acceleration applied to the sample, typically measured in comparison to the g. If samples are centrifuged long enough, the particles in the vessel will reach equilibrium wherein the particles accumulate specifically at a point in the vessel where their buoyant density is balanced with centrifugal force. Such an “equilibrium” centrifugation can allow extensive purification of a given particle. In sucrose gradient centrifugation, a linear concentration gradient of sugar (typically sucrose, glycerol, or a silica based density gradient media, like Percoll) is generated in a tube such that the highest concentration is on the bottom and lowest on top. Percoll is a trademark owned by GE Healthcare companies. A protein sample is then layered on top of the gradient and spun at high speeds in an ultracentrifuge. This causes heavy macromolecules to migrate towards the bottom of the tube faster than lighter material. During centrifugation in the absence of sucrose, as particles move farther and farther from the center of rotation, they experience more and more centrifugal force (the further they move, the faster they move). The problem with this is that the useful separation range of within the vessel is restricted to a small observable window. A properly designed sucrose gradient will counteract the increasing centrifugal force so the particles move in close proportion to the time they have been in the centrifugal field. Samples separated by these gradients are referred to as “rate zonal” centrifugations. After separating the protein/particles, the gradient is then fractionated and collected. Figure 3.3 Sucrose Density Gradient. Image derived from Michel Awkal back to the topPurification StrategyChoice of a starting material is key to the design of a purification process. In a plant or animal, a particular protein usually isn’t distributed homogeneously throughout the body; different organs or tissues have higher or lower concentrations of the protein. Use of only the tissues or organs with the highest concentration decreases the volumes needed to produce a given amount of purified protein. If the protein is present in low abundance, or if it has a high value, scientists may use recombinant DNA technology to develop cells that will produce large quantities of the desired protein (this is known as an expression system). Recombinant expression allows the protein to be tagged, e.g. by a His-tag or Strep-tag to facilitate purification, reducing the number of purification steps required. These techniques will be discussed in greater detail in Chapter 5. An analytical purification generally utilizes three properties to separate proteins. First, proteins may be purified according to their isoelectric points by running them through a pH graded gel or an ion exchange column. Second, proteins can be separated according to their size or molecular weight via size exclusion chromatography or by SDS-PAGE (sodium dodecyl sulfate-polyacrylamide gel electrophoresis) analysis. Proteins are often purified by using 2D-PAGE and are then analysed by peptide mass fingerprinting to establish the protein identity. This is very useful for scientific purposes and the detection limits for protein are nowadays very low and nanogram amounts of protein are sufficient for their analysis. Thirdly, proteins may be separated by polarity/hydrophobicity via high performance liquid chromatography or reversed-phase chromatography. Gel electrophoresis techniques are discussed in more detail in Section 3.2. This section will focus predominantly on chromatographic separations. For preparative protein purification, the purification protocol generally contains one or more chromatographic steps. The basic procedure in chromatography is to flow the solution containing the protein through a column packed with various materials. Different proteins interact differently with the column material, and can thus be separated by the time required to pass the column, or the conditions required to elute the protein from the column. Usually proteins are detected as they are coming off the column by their absorbance at 280 nm. Many different chromatographic methods exist, with the most common described below: Size Exclusion Chromatography (also known as Gel Filtration Chromatography)Chromatography can be used to separate protein in solution or under denaturing conditions by using porous gels. This technique is known as size exclusion chromatography. The principle is that smaller molecules have to traverse a larger volume in a porous matrix. Consequentially, proteins of a certain range in size will require a variable volume of eluent (solvent) before being collected at the other end of the column of gel. Thus, proteins will be separated based on their size (Figure 3.4). In the context of protein purification, the eluent is usually pooled in different test tubes. All test tubes containing no measurable trace of the protein to purify are discarded. The remaining solution is thus made of the protein to purify and any other similarly-sized proteins. Figure 3.4 Size Exclusion Chromatography. Also known as Gel Filtration Chromatography, is a low resolution isolation method that involves the use of beads that have tiny “tunnels” in them that each have a precise size. The size is referred to as an “exclusion limit,” which means that molecules above a certain molecular weight will not fit into the tunnels. Molecules with sizes larger than the exclusion limit do not enter the tunnels and pass through the column relatively quickly by making their way between the beads. Smaller molecules, which can enter the tunnels, do so, and thus, have a longer path that they take in passing through the column. Because of this, molecules larger than the exclusion limit will leave the column earlier, while smaller molecules that pass through the beads will elute from the column later. This method allows separation of molecules by their size. Image fromDr. Kevin Ahern and Indira Rajagopal Hydrophobic Interaction Chromatography (HIC)HIC media is amphiphilic, with both hydrophobic and hydrophilic regions, allowing for separation of proteins based on their surface hydrophobicity. Target proteins and their product aggregate species tend to have different hydrophobic properties and removing them via HIC further purifies the protein of interest. Additionally, the environment used typically employs less harsh denaturing conditions than other chromatography techniques, thus helping to preserve the protein of interest in its native and functional state. In pure water, the interactions between the resin and the hydrophobic regions of protein would be very weak, but this interaction is enhanced by applying a protein sample to HIC resin in high ionic strength buffer. The ionic strength of the buffer is then reduced to elute proteins in order of decreasing hydrophobicity (Figure 3.5). Figure 3.5 Hydrophobic Interaction Chromatography. The column matrix, shown in blue has a hydrophobic ligand covalently attached. In high salt conditions, proteins will bind to the matrix with differing affinity, with more hydrophobic proteins (shown in yellow) binding more tightly than more hydrophilic proteins (shown in green) When the salt concentration is decreased, proteins that are more hydrophilic will be released first, followed more hydrophobic proteins. Ion Exchange ChromatographyIon exchange chromatography separates compounds according to the nature and degree of their ionic charge. The column to be used is selected according to its type and strength of charge. Anion exchange resins have a positive charge and are used to retain and separate negatively charged compounds (anions), while cation exchange resins have a negative charge and are used to separate positively charged molecules (cations). Before the separation begins a buffer is pumped through the column to equilibrate the opposing charged ions. Upon injection of the sample, solute molecules will exchange with the buffer ions as each competes for the binding sites on the resin. The length of retention for each solute depends upon the strength of its charge. The most weakly charged compounds will elute first, followed by those with successively stronger charges. Because of the nature of the separating mechanism, pH, buffer type, buffer concentration, and temperature all play important roles in controlling the separation. Figure 3.6 demonstrates a type of ion exchange column known as a cation exchange column. In this case, the support consists of tiny beads to which are attached chemicals possessing a charge. Each charged molecule has a counter-ion. The figure shows the beads (blue) with negatively charged groups (red) attached. In this example, the counter-ion is sodium, which is positively charged. The negatively charged groups are unable to leave the beads, due to their covalent attachment, but the counter- ions can be “exchanged” for molecules of the same charge. Thus, a cation exchange columnwill have positively charged counter-ions and positively charged compounds present in a mixture passed through the column will exchange with the counter-ions and “stick” to the negatively charged groups on the beads. Molecules in the sample that are neutral or negatively charged will pass quickly through the column. On the other hand, in anion exchange chromatography, the chemical groups attached to the beads are positively charged and the counter-ions are negatively charged. Molecules in the sample that are negatively charged will “stick” and other molecules will pass through quickly. To remove the molecules “stuck” to a column, one simply needs to add a high concentration of the appropriate counter-ions to displace and release them. This method allows the recovery of all components of the mixture that share the same charge. Ion exchange chromatography is a very powerful tool for use in protein purification and is frequently used in both analytical and preparative separations. Figure 3.6 Cation Exchange Chromatography. In this diagram the negatively charged molecules (shown in red) are covalently attached to the column matrix beads (shown in blue). Sodium ions (Na+) are the counter ions that are replaced by positively charged proteins within the protein mixture. Neutral and negatively charged proteins do not stick and will pass through the column. The positively charged proteins can then be eluted from the column by adding higher concentrations of the counter ion (in this case the sodium ions). Image from Kevin Ahern and Indira Rajagopal Affinity Chromatography is a separation technique based upon molecular conformation, which frequently utilizes application specific resins. These resins have ligands (small molecules) attached to their surfaces which are specific for and will bind with the compounds to be separated. Most frequently, these ligands function in a fashion similar to that of antibody-antigen interactions. This “lock and key” fit between the ligand and its target compound makes it highly specific, frequently generating a single peak, while all else in the sample is unretained (Figure 3.7). For example, many membrane proteins are glycoproteins and can be purified by lectin affinity chromatography. Detergent-solubilized proteins can be allowed to bind to a chromatography resin that has been modified to have a covalently attached lectin. Proteins that do not bind to the lectin are washed away and then specifically bound glycoproteins can be eluted by adding a high concentration of a sugar that competes with the bound glycoproteins at the lectin binding site. Some lectins have high affinity binding to oligosaccharides of glycoproteins that is hard to compete with sugars, and bound glycoproteins need to be released by denaturing the lectin. Figure 3.7 Example of Affinity Chromatography. In this example, protein P1 has affinity for ligand Z and will bind to the column while proteins P2 and P3 will pass through the column. Protein P1 can then be eluted from the column using high concentrations of free ligand Z. A common technique involves engineering a sequence of 6 to 8 histidine residues into the N- or C-terminal of a recombinant protein. The polyhistidine binds strongly to divalent metal ions such as nickel and cobalt. The protein can be passed through a column containing immobilized nickel ions, which binds the polyhistidine tag. All untagged proteins pass through the column. The protein can be eluted with imidazole, which competes with the polyhistidine tag for binding to the column, or by a decrease in pH (typically to 4.5), which decreases the affinity of the tag for the resin. While this procedure is generally used for the purification of recombinant proteins with an engineered affinity tag (such as a 6xHis tag), it can also be used for natural proteins with an inherent affinity for divalent cations. Immunoaffinity chromatographyA special type of affinity chromatography is Immunoaffinity chromatography (Figure 3.8). This technique uses the specific binding of an antibody with its antigen (target molecule that the antibody will bind with selectively) to purify the protein of interest. The procedure involves immobilizing an antibody to a solid substrate (e.g. a porous bead or a membrane), which then selectively binds the target, while everything else flows through. The target protein can be eluted by changing the pH or the salinity. The immobilized ligand can be an antibody (such as Immunoglobulin G) or it can be a protein (such as Protein A). Because this method does not involve engineering in a tag, it can be used for proteins from natural sources. Antibody structure and their use in protein identification will be discussed in greater detail in Section 3.2. Figure 3.8. An Antigen Immunoprecipitation Experiment. The antibody is either pre-immobilized to a solid support (left) or immobilized using antibody binding proteins after incubation with the sample (right). Immobilization allows the immune complex to be extracted from the complex sample, washed and eluted providing a high enrichment of the protein under investigation Image from The Human Atlas Project back to the topHigh Performance Liquid Chromatography (HPLC) and Fast Protein Liquid Chromatography (FPLC)High performance liquid chromatography or high pressure liquid chromatography (HPLC)is a form of chromatography applying high pressure to drive the solutes through the column faster. This means that the diffusion is limited and the resolution is improved. The most common form is “reversed phase” HPLC, where the column material is hydrophobic. The proteins are eluted by a gradient of water and increasing amounts of an organic solvent, such as acetonitrile. The proteins elute according to their hydrophobicity. After purification by HPLC the protein is in a solution that only contains volatile compounds, and can easily be lyophilized (freeze dried). HPLC purification frequently results in denaturation of the purified proteins and is thus not applicable to proteins that do not spontaneously refold. Due to the drawbacks of HPLC, an alternative technique using a lower pressure system was developed and is called Fast protein liquid chromatography (FPLC). FPLC is a form of liquid chromatography that is often used to analyze or purify mixtures of proteins. As in other forms of chromatography, separation is possible because the different components of a mixture have different affinities for two materials, a moving fluid (the “mobile phase”) and a porous solid (the stationary phase). In FPLC the mobile phase is an aqueous solution, or “buffer”. The buffer flow rate is controlled by a positive-displacement pump and is normally kept constant, while the composition of the buffer can be varied by drawing fluids in different proportions from two or more external reservoirs. The stationary phase is a resin composed of beads, usually of cross-linked agarose, packed into a cylindrical glass or plastic column. FPLC resins are available in a wide range of bead sizes and surface ligands depending on the application. In the most common FPLC strategy, an ion exchange resin is typically chosen (Figure 3.9). A mixture containing one or more proteins of interest is dissolved in 100% buffer A and pumped into the column. The proteins of interest bind to the resin while other components are carried out in the buffer. The total flow rate of the buffer is kept constant; however, the proportion of Buffer B (the “elution” buffer) is gradually increased from 0% to 100% according to a programmed change in concentration (the “gradient”). Buffer B contains high concentrations of the exchanger ion. Thus as the concentration of the Buffer B gradually increases, bound proteins will dissociate depending on their ionic interactions with the column matrix and appear in the eluant. The eluant passes through two detectors which measure salt concentration (by conductivity) and protein concentration (by absorption of ultraviolet light at a wavelength of 280nm). As each protein is eluted it appears in the eluant as a “peak” in protein concentration and can be collected for further use. FPLC was developed and marketed in Sweden by Pharmacia in 1982 and was originally called fast performance liquid chromatography to contrast it with HPLC or high-performance liquid chromatography. FPLC is generally
applied only to proteins; however, because of the wide choice of resins and buffers it has broad applications. In contrast to HPLC the buffer pressure used is relatively low, typically less than 5 bar, but the flow rate is relatively high, typically 1-5 ml/min. FPLC can be readily scaled from analysis of milligrams of mixtures in columns with a total volume of 5 ml or less to industrial production of kilograms of purified protein in columns with volumes of many liters. Figure 3.9 Typical FPLC System. A. Scheme of basic compounents and typical flow path for a chromatography system. B. Picrue of GE Healthcare AKTA FPLC apparatus. Image provided by LaVerde, V., Dominici, P. and Astegno, A. (2017) Bio-protocol 7(8): e2230. Purification SchemeDuring the protein purification process it is necessary to have a quantitative system to determine how much protein has been purified, what concentration the protein represents from the original mixture, how biologically active the purified protein is, and the overall purity of the protein. This will help guide and optimize the purification method being developed. Ineffective separation techniques can be disregarded and other techniques that give higher yield or that retain biologically activity of the protein can be adopted. Thus, each step in the purification scheme is quantitatively evaluated for the following parameters: total protein, total activity, specific activity, yield, purification level. Each of these parameters will be defined within the sample protocol given below. Pretend you are a researcher that wants to isolate a novel, unknown protein from a bacterial culture. You grow 500 ml of the bacteria overnight at 37oC and harvest the bacteria by centrifugation. You remove the culture broth and retain the bacterial pellet. You then lyse the bacteria using freeze/thaw in 10 mL of reaction buffer. You then centrifuge the lysed bacteria to remove the insoluble materials and retain the supernatent that contains the soluble proteins. Your protein of interest has a biological activity that you can measure using a simple assay that causes a color change in the reaction mixture (Figure 3.10). You also note that this reaction rate increases with increasing concentrations of your protein supernatent (Figure 3.10) Figure 3.10. Example of a Chemical Reaction that causes a color change from orange to brown depending on increasing concentration. Image from: Ludwig, N., et. al. (2015) on Research Gate At this point, you can measure your baseline concentrations for the first purification level (bacterial lysis and removal of insoluble proteins and other cellular debris by centrifugation). Total Protein is calculated by measuring the concentration in a fraction of your sample, and then multiplying that by the total volume of your sample. In this case, you are starting with 10 mL of supernatent. In a typical assay to measure protein concentration, you will use 50 – 200 μL of sample to determine the protein concentration. For example, if you calculate that there is 7.5 μg/μL in your initial assay, you would need to convert that value into mg/mL and then multiply it by 10 mL for a total of 75 mg of protein in 10 mL of supernatant (Table 3.1) Total Activityis measured as the enzyme activity within the assay, multiplied by the total volume of the sample. For example, in the initial sample, you might use 5 to 50 μL of sample in your biological reaction (Figure 3.10). If you calculated the activity in your assay to be 2.5 units/μL, this would be equivalent to 2,500 units/mL or 25,000 units/10 mL of supernatant. Note that, the enzyme unit, or international unit for enzyme (symbol U, sometimes also IU) is a unit of enzyme’s catalytic activity. 1 U (μmol/min) is defined as the amount of the enzyme that catalyzes the conversion of one micromole of substrate per minute under the specified conditions of the assay method. Specific Activity is measured by dividing the Total Activity by the Total Protein. In our example, 25,000 units divided by 75 mg of protein = 333.3 units/mg. Yield is a measure of the biological activity retained in the sample after each purification step. The amount in the first step is set to be 100%. All subsequent yield steps will be evaluated using the first purification step. It is calculated by dividing the total activity of the current step, by the total activity of the first step and then multiplying by 100. Purification levelevaluates the purity of the protein of interest by dividing the specific activity calculated after each purification step by the specific activity of the first purification step. Thus, the first step always has a value of 1. Table 3.1 Typical Protein Purification Scheme Note that after each purification step that the Total Protein goes down, as you are purifying your protein away from other proteins in the mixture. Total Activity also goes down with each purification step, as some of your protein of interest is also lost at each purification step, because (1) some protein will stick to the test tubes and glassware, (2) some protein won’t bind with 100% efficiency to your column matrix, (3) some protein may bind too tightly to be removed from the column matrix during elution, and (4) some protein may be denatured or degraded during the purification process. The amount of your protein of interest that is lost is represented within the overall percent yieldfor each purification step. If the percent yield is too low alternative purification methods should be explored. Note that in a good protein purification scheme that the specific activity should go up substantially with each level
of purification as the amount of your protein of interest makes up a greater percentage of the total protein within that fraction. If the specific activity only increases modestly within a purification step, or if it decreases during a purification step, this could indicate that (1) your protein of interest is being substantially lost at that step, (2) that your protein of interest is being denatured or degraded and is no longer biologically active, or (3) that a required cofactor or binding
protein is being reduced at that purification step. Additional experiments may need to be conducted to determine which of the causes predominates, so that steps can be taken to reduce protein inactivation. For example, many proteins are temperature sensitive and will degrade or denature at room temperature. Completing purification steps on ice can often reduce degradation. Overall, the fold increase in purification level should increase exponentially during the purification process. Note that in our example, if after 4 steps of purification our proteins is close to 95% pure, this would indicate that our protein of interest makes up approximately 1.24% of the total protein within the sample. back to the top3.2 Protein Identification and VisualizationAnalytical techniques that can be used to positively identify or visualize a protein of interest within a mixture can also be a valuable tool to understanding the biological activity and significance of a protein within a living system and can also be used to help guide protein purification schemes. Gel Electrophoresis(work derived from Magdeldin, S.) Agarose is a natural linear polymer extracted from seaweed that forms a gel matrix by hydrogen-bonding when heated in a buffer and allowed to cool. For most applications, only a single-component agarose is needed and no polymerization catalysts are required (Figure 3.11). Therefore, agarose gels are simple and rapid to prepare. They are the most popular medium for the separation of moderate and large-sized nucleic acids and have a wide range of separation but a relatively low resolving power, since the bands formed in the gels tend to be fuzzy and spread apart. This is a result of pore size and cannot be largely controlled. These and other advantages and disadvantages of using agarose gels for electrophoresis are summarized in Table 3.2. Agarose gels are not typically used for protein samples and won’t be discussed in this chapter further. However, they will be revisted in Chapter 5 covering nucleic acid techniques. Table 3.2. Advantages and Disadvantages of Agarose Gel Electrophoresis. Polyacrylamide gels are chemically cross-linked gels formed by the polymerization of acrylamide with a cross-linking agent, usually N,N’-methylenebisacrylamide (Figure 3.11). The reaction is a free radical polymerization, usually carried out with ammonium persulfate as the initiator and N,N,N’,N’-tetramethylethylendiamine (TEMED) as the catalyst. Although the gels are generally more difficult to prepare and handle, involving a longer time for preparation than agarose gels, they have major advantages over agarose gels. They have a greater resolving power, can accommodate larger quantities of sample without significant loss in resolution and the purity of the sample recovered from polyacrylamide gels is extremely high. Moreover, the pore size of the polyacrylamide gels can be altered in an easy and controllable fashion by changing the concentrations of the two monomers. Thus, it is commonly used to separate proteins and smaller fragments of DNA. It should be noted that polyacrylamide is a neurotoxin (when unpolymerized), but with proper laboratory care it is no more dangerous than various commonly used chemicals. Some advantages and disadvantages of using polyacrylamide gels for electrophoresis are depicted in Table 3.3. Table 3.3. Advantages and Disadvantages of Polyacrylamide Gel Electrophoresis. Hydrated gel networks have many desirable properties for electrophoresis. They allow a wide variety of mechanically stable experimental formats such as horizontal/vertical electrophoresis in slab gels or electrophoresis in tubes or capillaries. The mechanical stability also facilitates post electrophoretic manipulation making further experimentation possible such as blotting, electro-elution or mass spectral identification /finger printing of intact proteins or of proteins digested in gel slices. Since gels used in biochemistry are chemically rather unreactive, they interact minimally with biomolecules during electrophoresis allowing separation based on physical rather than chemical differences between sample components. Figure 3.11 Gels Commonly Used in Electrophoresis. (A) Agarose is composed of agarbiose, (B) The polymerization of acrylamide and bisacrylamide to form polyacrylamide gel. The polymerization reaction is initiated by persulfate radicals and catalyzed by TEMED. Image from Magdeldin, S. Gel electrophoresis of proteins with a polyacrylamide matrix, commonly called polyacrylamide gel electrophoresis (PAGE)is undoubtedly one of the most widely used techniques to characterize complex protein mixtures. It is a convenient, fast and inexpensive method because they require only the order of micrograms quantities of protein. They are usually run in a vertical format and the gel rigs contain an upper and lower buffer reservoir (Fig. 3.12A). The samples are loaded in wells that contact the upper buffer reservoir which will house the negative cathode. The proteins migrate towards the positive anode when the electric current is applied. Note that proteins have a net electrical charge if they are in a medium having a pH different from their isoelectric point and therefore have the ability to move when subjected to an electric field. The migration velocity is proportional to the ratio between the charges of the protein and its mass. The higher charge per unit of mass the faster the migration. Proteins do not have a predictable structure as nucleic acids, and thus their rates of migration are not similar to each other. Furthermore, they will not migrate when applying an electromotive force, when the pH of the system is the same as isoelectric point. PAGE gels that are run in this fashion are called Native PAGE, as the proteins are still folded in their native state found in vivo. In this situation, proteins migrate according to their charge, size and shape. Alternatively, proteins may be denatured prior to electrophoresis. The most common way to denature the proteins is by adding a detergent such as sodium dodecyl sulfate (SDS) (Fig 3.12B). This not only denatures the proteins, but it also coats the protein with a negative charge, such that all of the proteins will run towards the positive lead when placed into an electric field. This type of electrophoresis is referred to as SDS-PAGE and separates proteins exclusively according to molecular weight. A reducing agent that breaks disulfide bonds, such as dithiothreitol (DTT) is often added to the loading buffer as well, causing proteins to fully denature and dissociate into the monomer subunits (Fig 3.12C). This ensures that the proteins migrate through the gel in direct relation to their size, rather than by charge or shape. Figure 3.12 Polyacrylamide Gel Electrophoresis (PAGE). (A) Shows a typical set up for PAGE. A vertical gel is placed into a rig with an upper and lower buffer reservoir. The upper reservoir contains the gel wells where the protein is loaded and will house the cathode (negative charge). The proteins will run towards the anode (positive charge) when an electric field is placed on the system, in relation to the protein size, shape and charge. (B) Sodium dodecyl sulfate (SDS) is often used to denature proteins prior to PAGE analysis, causing proteins to migrate based on size only. (C) Reducing agents, such as dithiothreitol (DTT) are often used in combination with SDS, to ensure that disulfide bonds within the protein or between protein subunits are fully reduced to free cysteine residues. Figures from: (A) Bensaccount (B) Fdardel, and (C) Edgar181 Detection of Proteins in GelsProteins separated on a polyacrylamide gel can be detected by various methods, for instance dyes and silver staining (Figure 3.13).
The Coomassie blue staining allows detecting up to 0.2 to 0.6 µg of protein, and is quantitative (linear) up to 15 to 20 µg. It is often used in methanol-acetic acid solutions and is discolored in isopropanol-acetic acid solutions (Fig. 1 A). For staining of 2-DE gels it is recommended to remove ampholytes by adding trichloroacetic (TCA) to the dye and subsequently discolor with acetic acid.
It is an alternative to routine staining protein gels (as well as nucleic acids and lipopolysaccharides) because its ease use and high sensitivity (50 to 100 times more sensitive than Coomassie blue staining) (Fig. 1 B). This staining technique is particularly suitable for two-dimensional gels.
The autoradiography is a detection technique of radioactively labeled molecules that uses photographic emulsions sensitive to radioactive particles or light produced by an intermediate molecule. The emulsion containing silver is sensitive to particulate radiation (alpha, beta) or electromagnetic radiation (gamma, light…), so that it precipitates as metallic silver. The emulsion will develop as dark precipitates in the region in which radioactive proteins are detected. Figure 3.13. SDS-PAGE. Proteins separated on SDS-PAGE and detected by Coomassie blue (A) and silver staining (B). Standards of proteins to know molecular weight are also loaded at edges. Image from Magdeldin, S. Isoelectric FocusingThis technique is based on the movement of molecules in a pH gradient. Amphoteric molecules such as amino acids and proteins are separated in an environment where there is a difference of potential and pH gradient. The region of the anode (+) is acidic and the cathode (-) is alkaline. Between them down a pH gradient such that the molecules to be separated have their isoelectric point within the range. Substances that are initially in regions with a pH below its isoelectric point are positively charged and migrate towards the cathode, while those that are in media with pH lower than its pI will have negative charge and migrate towards the anode (Figure 3.14). The migration will lead to a region where the pH coincide with its pI, have a zero net charge (form zwitterions) and stop. Thus amphoteric molecules are located in narrow bands where the pI coincides with the pH. In this technique the point of application is not critical as molecules will always move to their pI region. The stable pH gradient between the electrodes is achieved using a mixture of low molecular weight ampholytes which pI covers a preset range of pH. Figure 3.14. Isoelectric Focusing. A pH gradient is established in a gel before loading the sample. After the sample is loaded a voltage is applied. The protein will migrate to their isoelectric pH, which they have no net charge. Image from Magdeldin, S. Two-Dimensional Gel Electrophoresis |